Electro-mechanically stretched micro fibers and methods of use thereof

ABSTRACT

The presently disclosed subject matter provides a scalable and electrostretching approach for generating hydrogel microfibers exhibiting uniaxial alignment from aqueous polymer solutions. Such hydrogel microfibers can be generated from a variety of water-soluble natural polymers or synthetic polymers. The hydrogel microfibers can be used for controlled release of bioactive agents. The internal uniaxial alignment exhibited by the presently disclosed hydrogel fibers provides improved mechanical properties to hydrogel microfibers, and contact guidance cues and induces alignment for cells seeded on or within the hydrogel microfibers.

CROSS-REFERENCE TO RELATED APPLICATIONS

This application is a Continuation in Part (CIP) of U.S. applicationSer. No. 14/398,072 that claims the benefit of International PatentApplication No. PCT/US2013/038805, filed Apr. 30, 2013, and U.S.Provisional Application Nos. 61/640,057, filed Apr. 30, 2012, and61/665,498, filed Jun. 28, 2012; and each of which is incorporatedherein by reference in its entirety.

FEDERALLY SPONSORED RESEARCH OR DEVELOPMENT

This invention was made with government support under DMR-0748340awarded by the National Science Foundation. The government has certainrights in the invention.

BACKGROUND

Hydrogels have been widely investigated for a variety of biomedicalapplications, particularly as scaffolds offering a 3-dimensional (3D)microenvironment for tissue regeneration. Hydrogels have been used for3D cell culture and tissue regeneration because of their high watercontent resembling the aqueous microenvironment of the naturalextracellular matrix (Seliktar, 2012) and tunable biochemical andphysicochemical properties (Burdick and Anseth, 2002; Williams et al.,2003; Silva et al., 2004). While many properties of natural hydrogelmatrices are modifiable, their inherent isotropic structure limits thecontrol over cellular organization that is critical to restore tissuefunction.

Previous studies have primarily focused on exploring the mechanical andbiochemical versatility of hydrogels and elucidating their impact oncellular activities (Engler et al., 2006; Discher et al., 2005; Lutolfet al., 2003; Dalsin et al., 2003; Martino et al., 2009). A lack ofmethodologies exists, however, for engineering anisotropic topographicalcues in hydrogels to control the 3D spatial patterns of encapsulatedcells. As a result, controlling topographically induced cell alignmentand migration has not been readily achieved for hydrogel matrices, eventhough such cellular manipulation on 2D substrates has been shown to beimportant in controlling cell organization, tissue microarchitecture,and biological function (Yang et al., 2005; Bettinger et al., 2009; Chewet al., 2008; Aubin et al., 2010).

Recently, Kang et al. reported a microfluidic-based alginate hydrogelmicrofiber with a surface alignment feature produced by solutionextrusion through a grooved round channel, and demonstrated guidedneurite outgrowth for neurons cultured on the surface of the microfibers(Kang et al., 2011). This alignment cue, however, is only confined tothe surface of the microfibers. Zhang et al. have generated peptidenanofiber hydrogels with long-range nanofiber alignment throughheat-assisted self-assembly of amphiphilic peptide molecules andmechanical shear (Zhang et al., 2010). Although the resulting alignednanofiber “noodles” effectively induced cellular alignment in 3D, thismethod is only applicable to specific peptide materials.

On the other hand, cellular alignment mediated by 2D electrospunnanofiber matrices has been shown to effectively promote stem celldifferentiation and cellular functions (Lim and Mao, 2009; Ji et al.,2006). Although dispersing solid polymer nanofibers into the hydrogelmatrix has been used to generate a composite scaffold (Coburn et al.,2011), controlling alignment of the nanofibers inside a hydrogel matrixis challenging.

The development of artificial microvascular vessels is of criticalimportance in the fields of cardiovascular diseases, cancer growth, andtissue engineering of vascularized organs. In tissue engineering, thereis currently a limit on the size of tissues that can be constructed invitro due to the diffusion limit of nutrients into developing organs. Assuch, only thin tissues like skin grafts or avascular tissues likecartilage can be successfully engineered in vitro, with largerconstructs requiring the existence of a vascular network within itselfto supply the necessary nutrients for cell survival and to avoidnecrotic areas post-implantation (Huang, G. Y., et al., Biofabrication,2011. 3(1):012001, and Auger, F. A., et al., Annual Rev BiomedEngineering, 2013. 15(1):177-200. Development of functionalmicrovascular vessels requires complex interactions between endothelialcells (ECs), perivascular cells, and the extracellular matrix (ECM)(Jain R K, Nat Med, 2003; 9: 685-693; Carmeliet P and Jain R K, Nature,2000; 407: 249-257). The ECM is comprised of an abundance of nanometersized macromolecular ECM proteins. As such, many of the physicalinteractions between vascular cells and macromolecular components of theECM occur at the sub-micron scale. EC adhesion to the ECM initiates theangiogenic cascade (Hynes R O, J. Thromb Haemost, 2007; 5: 32-40). Cellsdetect and respond to the nanoarchitecture of their microenvironment bycytoskeletal reorganization and activated signaling cascades to regulatefundamental cell behaviors. Indeed, it has been shown that surfaces withnano-scale line-grating features affect EC adhesion, alignment, andelongation (Ranjan A, and Webster T, Nanotechnology, 2009; 20: 305102;Liliensiek S, et al., Biomaterials, 2010; 31: 5418-5426; Bettinger C J,et al., Adv Mater, 2008; 20: 99-103; Lu J, et al., Acta Biomater, 2008;4: 192-201).

Arterioles are the blood vessels found immediately before capillaries,ranging from tens to hundreds of microns in diameter. ECs, sitting ontheir basal lamina, comprise the innermost lining of the vessels, thislayer containing mainly collagen type IV (Col IV), fibronectin (Fn),laminin (Lmn), and heparin sulfate proteoglycan (Roy, S., et al.,Current Eye Research, 2010; 35(12):1045-1056). This is followed by alayer of subendothelial connective tissue and an internal elastic lamina(Hibbs, R. G., et al., Am Heart J, 1958; 56(5):662-670; Yen, A. and I.M. Braverman, J Investig Dermatol, 1976; 66(3):131-142; Weber, K. and O.Braun-Falco, Archiv für dermatologische Forschung, 1973; 248(1):29-44),which are composed of different ECM proteins, mainly of collagens suchas type I and III (Col I, Col III), and elastin (Eln) (Yen, A. JInvestig Dermatol, 1976; Tsamis, A., et al., J Royal Society Interface,2013; 10(83):20121004. These layers together make the tunica intima ofblood vessels. Mural cells, along with their ECM, make up the tunicamedia (or middle layer), a layer of perivascular cells increasing inthickness with vessel size (Yen, A. J Investig Dermatol, 1976; Marieb,E. N. and K. Hoehn, Human anatomy & physiology. 2007: Pearson Education;Standring, S., Gray's anatomy. The anatomical basis of clinicalpractice, 2008. 39), and provide the contractility necessary forvasoreactivity. In the smallest of arterioles, the perivascular cellsare pericytes, which increase in confluency with vessels size andeventually are replaced by vascular smooth muscle cells (vSMCs) (Yen, A.J Investig Dermatol, 1976; Weber, K., 1973; Marieb, E. N. and K. Hoehn,2007: Pearson Education; Standring, S., Gray's anatomy, 2008:39). Thetunica media in arterioles has a primarily circumferential orientationof the vSMCs, which is necessary for vasoconstriction. Opposite toarterioles, venules collect blood from the capillary beds and are alsosupported by perivascular cells, though here the tunica media does notfollow a circumferential organization and the ECM layers are less robustthan in arterioles ((Hibbs, R. G., et al., Am Heart J, 1958; Yen, A. andI. M. Braverman, J Investig Dermatol, 1976; Marieb, E. N. and K. Hoehn,2007: Pearson Education; Standring, S., Gray's anatomy, 2008:39; Jain,R. K., Nature medicine, 2003; 9(6): p. 685-693). On top of the tunicamedia layer lays the tunica adventitia, though it is only present inlarger blood vessels (Carmeliet P and Jain R K, Nature, 2000; 407:249-257; Bruce Alberts A J, et al., Molecular Biology of the Cell, 2002,New York: Garland Science; Jain R K, Nature Medicine, 2003; 9: 685-693;Schwartz S M, and Benditt E P, American J Pathology, 1972; 66: 241;Schriefl A J, et al., Journal of The Royal Society Interface, 2012; 9:1275-1286; Tsamis A, et al., Journal of The Royal Society Interface;2013; 10; Canham, P B, et al., Cardiovascular 1989; 23: 973-982; FinlayH, et al., Journal of vascular research; 1995; 32: 301-312; Movat H Z,et al., Experimental and Molecular Pathology, 1963; 2: 549-563). Thespecific composition as well as the organization and arrangement of bothcellular and ECM components in each layer are necessary for propermicrovasculature development, maturation, stability, and function (JainR K, Nat Med, 2003; 9: 685-693; Carmeliet P and Jain R K, Nature, 2000;407: 249-257).

It has been shown that in microvasculature EC nuclei and cytoskeletonare aligned in the direction of blood flow. However, studies onmicrovasculature have not analyzed the organization of different ECMcomponents in detail (Movat H Z, and Fernando N V P, Experimental andMolecular Pathology, 1963; 2: 549-563; Hibbs R G, et al., American HeartJournal, 1958; 56: 662-670; Fernando N V, and Movat H Z, Experimentaland Molecular Pathology, 1964; 3: 1-9). The literature that has studiedthe particular organization of the ECM in native blood vessels isfocused on analyzing the elastin and collagen structures in the aortaand other large arteries (Schwartz S M, and Benditt E P, American JPatholo, 1972; 66: 241; Schriefl A J, et al., Journal of The RoyalSociety Interface, 2012; 9: 1275-1286; Tsamis A, et al., Journal of TheRoyal Society Interface; 2013; 10; Canham, P B, et al., Cardiovascular,1989; 23: 973-982; Finlay H, et al., Journal of vascular research, 1995;32: 301-312; Halloran B G, et al., Journal of Surgical Research, 1995;59: 17-22). These studies have shown that the structural organization ofcollagen varies not only between the three layers of the vasculature,but also varies with vessel size and specific location in the body(Tsamis A, et al., Journal of The Royal Society Interface, 2013; 10).

Overall, the studies agree that each tunica possesses at least twodifferent families of collagen fibrils, with distinctly differentorganizations. The general arrangement has been found to be close toaxial in the adventitia, outer layers having a more pronounced axialorientation transitioning to a circumferential alignment in inner layers(Finlay H, et al., Journal of vascular research, 1995; 32: 301-312). Incontrast, the medial layer has been shown to have a nearly perfectcircumferential order (Schriefl A J, et al., Journal of The RoyalSociety Interface, 2012; 9: 1275-1286; Tsamis A, et al., Journal of TheRoyal Society Interface; 2013; 10; Canham, P B, et al., Cardiovascular,1989; 23: 973-982; Finlay H, et al., Journal of vascular research, 1995;32: 301-312). Studies also agree both the internal and external elasticlamina are fenestrated, though both axial and circumferentialorganization of these layers has been reported (Movat H Z, and FernandoN V P, Experimental and Molecular Pathology, 1963; 2: 549-563; Moore DH, and Ruska H, J Biophysical and Biochemical Cytology, 1957; 3:457-462). On the other hand, the subendothelium has been found to have amore varied composition. It has been described as a multilayered fabricof collagen, containing distinct layers of both longitudinally andcircumferentially aligned fibers (Schwartz S M, and Benditt E P,American J Pathology, 1972; 66: 241; Canham, P B, et al.,Cardiovascular, 1989; 23: 973-982; Finlay H, et al., Journal of vascularresearch, 1995; 32: 301-312; Gasser T C, et al., J Royal SocietyInterface, 2006; 3: 15-35); some studies suggesting a layer oflongitudinally aligned ECM directly under the media layer, followed by ahelically arranged region of connective tissue beneath, and a thincircumferentially aligned layer next to the lumen (Finlay H, et al.,Journal of vascular research, 1995; 32: 301-312). However, thesefindings were all made on large arteries with diameters in themillimeter range.

The creation of clinically relevant functional microvascular conduits(i.e. arterioles and venules) remains a challenge. To date, only a fewstudies have attempted to model or recreate microvasculature in a full3D setting in vitro (Miller J S, et al., Nat Mater, 2012; 11: 768-774;Zheng Y, et al., PNAS USA, 2012; 109: 9342-9347; Neumann T, et al.,Microvascular Research, 2003; 66: 59-67; Norotte, C., et al.,Biomaterials, 2009; 30 (30): p. 5910-5917), though none has achieved afully functional microvascular structure recapitulating both thecellular and ECM protein organization in the multilayer formationpresent in native vasculature. The majority of models have eitherfocused on the development of capillary beds in natural or synthetichydrogels (Yee D, et al., Tissue Engineering Part A, 2011; 1351-1361;Hanjaya-Putra D, et al., J Cellular Molecular Medicine, 2010; 14:2436-2447; Meyer G T, et al., Anat. Rec., 1997; 249: 327-340; Bayless KJ, Davis G E, J Cell Science, 2002; 115: 1123-1136; Bayless K J,American J Pathology, 2000; 156: 1673-1683; Grant D S, et al., CellPress, 1989; 933-943; Sacharidou A, et al., Cells Tissues Organs, 2011;195: 122-143; Davis G E, and Camarillo C W, Experimental Cell Research1996; 224: 39-51; Kim D J, et al., Blood, 2013; 121(17):3521-30;Hanjaya-Putra D, et al., Blood, 2011; 118: 804-815; Lutolf M, andHubbell J, Nature biotechnology, 2005; 23: 47-55; Moon J J, et al.,Biomaterials, 2010; 31: 3840-3847), decellularized matrices (Hielscher AC, et al., American Journal of Physiology—Cell Physiology, 2012; 302:C1243-C1256; Soucy P A, and Romer L H, Matrix Biology, 2009; 28:273-283), and electrospun polymer scaffolds (Pham Q P, et al., TissueEng, 2006; 12: 1197-1211; Kumbar S G, et al., Biomed Mater, 2008; 3:034002), or have instead aimed to create vascular grafts typically >3 mmin diameter (Aper T, Schmidt et al., European Journal of Vascular andEndovascular Surgery, 2007; 33: 33-39; Gui L, Tissue engineering Part A,2009; 15: 2665-2676; Swartz D D, et al., American Journal ofPhysiology-Heart and Circulatory Physiology, 2005; 288: H1451-H1460;Kaushal S, et al., Nature medicine, 2001; 7: 1035-1040; Cho S-W, et al.,Annals of surgery, 2005; 241: 506; L'heureux N, et al., The FASEBJournal, 1998; 12: 47-56). While these models enabled study of theECM-driven molecular mechanisms that regulate EC tubulogenesis, theymostly support spontaneous and random tubulogenesis (size, shape,organization, etc.). Recent work employs micro-patterning anddemonstrates an organized vascular network structure within hydrogels,some of which are able to recruit vascular smooth muscle cells (vSMCs)(Baranski J D, et al., Proc Natl Acad Sci USA, 2013; Miller J S, et al.,Nat Mater, 2012; 11: 768-774; Zheng Y, et al., Proc Natl Acad Sci USA,2012; 109: 9342-9347). Nonetheless, these systems have limited controlover topographical cues presented by the ECM and impart a barrier forthe high-resolution, dynamic, and detailed study of vascularorganization as well as specific cell-ECM and multi-cellularinteractions.

Tubular polymeric scaffolds have the potential to provide a better andmore sophisticated platform to study the microvasculature, but currentlyare obtainable with diameters in the millimeter range (Melchiorri A J,et al., Tissue Eng Part B Rev, 2012; Fioretta E S, et al., MacromolBiosci, 2012; 12: 577-590; Gui L, et al., Tissue Eng Part A, 2009; 15:2665-2676) and are mostly used to study graft's mechanical strength (WuW, et al., Nat Med, 2012; 18: 1148-1153; Huynh T, and Tranquillo R,Annals of Biomedical Engineering, 2010; 38: 2226-2236; Lee K-W, et al.,PNAS, 2011; 108: 2705-2710). To recapitulate the microvasculature invitro, which has not previously been obtainable, the tubular scaffoldsmust exhibit a physiologically-relevant diameter and sub-microntopography with sufficient mechanical properties, be biocompatible,mediate specific cell adhesion, allow tubular vessel formation, and,support multi-cellular interactions, thus generating microvessels thatmimic natural structure and properties.

Endothelial colony forming cells (ECFCs), a subpopulation of endothelialprogenitor cells, are known for their proliferative capacity andcontribution to functional vessels (Critser P J, and Yoder M C, CurrOpin Organ Transplant, 2010; 15: 68-72; Yoder M C, J of Thrombosis andHaemostasis, 2009; 7: 49-52; Yoder M C, et al., Blood, 2007; 109:1801-1809). Recently, we revealed that ECFCs deposit ECM proteins,namely collagen IV, fibronectin and laminin, and also assemble ECM intoweb-like structures when cultured on Petri dishes (Kusuma S, et al.,FASEB J, 2012; 26: 4925-4936). This finding suggests an important rolefor ECM production by ECFCs in the process of vascular assembly that hasnot yet been identified.

The cellular and ECM composition, as well as their specific structuralorganization, varies greatly in blood vessels of different type, size,and function. While postcapillary venules (10-30 μm) are formed by ECsand their basal lamina, along with scattered pericytes, and aresemipermeable like capillaries (Standring S, 2008, Gray's anatomy),larger venules (larger than 50 μm in diameter) have a muscle layer and athin adventitia (Standring S, 2008). Both arterioles of 100 μm to 300 μmin diameter and muscular arteries (300 μm to 1 cm) have an alignedendothelium sitting on its basal lamina, surrounded by a layer ofconnective tissue. This layer is followed by a well-defined fenestratedinternal elastic membrane (with ECM occupying void spaces in thefenestrae) and a developed tunica media composed of several layers ofcircumferentially oriented vSMCs and ECM (mainly collagenous and elasticfibrils). These arteries are the main vessels in charge of restrictingblood flow to capillary beds via vSMC constriction in response to neuralor chemical stimuli (Standring S, 2008; Marieb E, Hoehn K, Human Anatomy& Physiology (8 ed., 2010), Boston, Mass.: Pearson Benjamin Cummings).Even though the ECM is known to be circumferentially oriented in thetunica media, the specific alignment of each layer in the tunica intimaremains unclear; several studies report a mix of circumferentialorientation close to the lumen and axial orientation close to the media(Schwartz S M, and Benditt E P, American J Pathology, 1972; 66: 241-264;Schriefl A J, et al., Journal of The Royal Society Interface, 2012; 9:1275-1286; Tsamis A, et al., Journal of The Royal Society Interface;2013; Movat H Z, et al., Experimental and Molecular Pathology, 1963; 2:549-563; Fernando N V, and Movat H Z, Experimental and MolecularPathology, 1964; 3: 1-9). However, it is known that circumferentiallyaligned ECM is necessary for vessels to be able to withstand thecircumferential stress resulting from the distending pressure of bloodflow. These facts highlight the importance of having bothcircumferential alignment of ECM and several layers of vSMCs, an aspectnot found in current microvasculature models. Furthermore, theorganization of other ECM components, such as fibronectin and laminin,remains widely uninvestigated. There is a need for a model to study themicrovasculature that recapitulates endothelial cell and perivascularcell alignment and ECM deposition for generating microvessels.

SUMMARY

In some aspects, the presently disclosed subject matter provides amethod for preparing a microfiber having a uniaxial alignment, themethod comprising: (a) providing at least one starting solutioncomprising one or more polymers; (b) applying an electrical potential tothe at least one starting solution sufficient to initiate a jet streamof polymer solution; and mechanically stretching the jet stream ofpolymer solution during or after collecting the jet stream of polymersolution in a collection bath comprising a stabilizing solution, whereinthe collection bath is positioned at a separation distance such that thejet stream of polymer solution is collected before it is accelerated byan electrical field created by the applied electrical potential.

In other aspects, the presently disclosed subject matter provides amicrofiber prepared by the presently disclosed methods.

Certain aspects of the presently disclosed subject matter having beenstated hereinabove, which are addressed in whole or in part by thepresently disclosed subject matter, other aspects will become evident asthe description proceeds when taken in connection with the accompanyingExamples and Figures as best described herein below.

Another embodiment of the present invention is a three-dimensional (3D)fibrin microfiber scaffold for a novel in vitro model of themicrovasculature that recapitulates endothelial cell alignment and ECMdeposition. The microfiber scaffold allows the sequential co-culture ofendothelial cells and other cells, such as perivascular cells. Thismodel establishes that the fiber curvature affects the circumferentialdeposition of ECM from endothelial cells independently of cellularorganization. The invention further presents a multicellularmicrovascular structure with an organized endothelium and multicellularperivascular tunica media. The present invention provides uniqueopportunities to assess microvasculature development and regeneration.

Another embodiment of the present invention relates to a tubularpolymeric scaffold, for example a polymer hydrogel microfiber, comprisedof electrostretched polymer nanofibers. The polymer preferably isalginate, fibrin (fibrinogen), gelatin, hyaluronic acid, or combinationsthereof. More preferably, the polymer is fibrin. An electrostretchedpolymer microfiber serves as a template for the step-wise creation ofmicrovasculature in vitro.

An embodiment of the invention relates to an electrostretched polymermicrofiber that has uniform and tunable diameter, while preserving analigned internal and external nanotopography. The polymer preferably isalginate, fibrin (fibrinogen), gelatin, hyaluronic acid, or combinationsthereof. In a particular embodiment, the polymer is fibrin. Preferably,the polymer microfiber has a diameter of about 500 μm or less, morepreferably from about 100 μm to about 450 μm.

In an embodiment of the invention, the electrostretched polymermicrofiber generates a micro-cylindrical fiber with a line gratingnanotopography. The microfiber enables both endothelial layerorganization and co-culture with a second cell type, for examplesupporting mural cells (perivascular cells).

In some embodiments, the invention relates to a polymer microfiberseeded with endothelial progenitor cells, such as an endothelial colonyforming cells. The endothelial cells adhere to the surface of the fibrinmicrofiber, and align longitudinally with the polymer microfiber. Theattached endothelial cells deposit extracellular matrix (ECM)circumferentially organized depending on the size of the microfiber. Theextracellular matrix is composed of proteins that encircle (wrap around)the microfiber along the fiber circumference, perpendicular to the cellorientation. In embodiments, the extracellular matrix proteins may belaminin, collagen IV, and/or fibronectin.

Other embodiments relate to endothelial colony forming cells on polymermicrofibers that deposit extracellular matrix proteins. Theextracellular matrix proteins include, for example, laminin, collagenIV, and fibronectin. The extracellular matrix proteins wrap around thepolymer microfiber, perpendicular to the cell orientation, along thefiber's circumference. Further, the extracellular matrix proteins can beabove, among, or below the endothelial cells (between the cells and themicrofiber).

Embodiments of the invention relate to the polymer microfibers culturedwith endothelial progenitor cells being further seeded with a secondcell type, such as mural cells (perivascular cells). Preferably, themural cells are vascular smooth muscle cells or pericytes. In oneembodiment, the vascular smooth muscle cells deposit ECM proteins, forexample collagen type I, collagen type III and/or elastin. Embodimentsinclude the ECM proteins, for example collagen types I and III, and.elastin, located beneath the vascular smooth muscle cell layer and abovethe endothelial progenitor cell layer. In another embodiment, pericytesdeposit ECM proteins, for example collagen types III and IV. Thecollagen type IV is located above the endothelial progenitor cell layer.

Other embodiments of the invention are microvascular structuresincluding an electrostretched polymer microfiber seeded with endothelialprogenitor cells. Endothelial progenitor cells, such as endothelialcolony forming cells, adhere to the surface of the polymer microfiber,align longitudinally with the microfiber, and deposit extracellularmatrix circumferentially organized. Extracellular matrix proteins,including for example, laminin, collagen IV, and fibronectin, canencircle the polymer microfiber. The extracellular matrix proteins areoriented perpendicular to the cell orientation, along the fiber'scircumference. Further, the extracellular matrix proteins are above,below, or among the endothelial cells. The term “below the endothelialcells” means that the ECM protein is between the cells and themicrofiber. In some embodiments the microvascular structure comprisingan electrostretched polymer microfiber seeded with endothelialprogenitor cells is further seeded with a second cell type, for examplemural cells. Preferably, the mural cells are vascular smooth musclecells or pericytes. In some embodiments, the vascular smooth musclecells deposits ECM proteins, for example collagens type I and type III,fibronectin, laminin, and elastin. Embodiments include the ECM proteins,for example collagen type I, type III and elastin located beneath thevascular smooth muscle cell layer and above the endothelial progenitorcell layer. In other embodiments, pericytes deposit ECM proteins, forexample collagen types I, III and IV, fibronectin and laminin. Thecollagen type IV is located above the endothelial progenitor cell layer.

In further embodiments of the invention, the polymer microfiber inducesincreased deposition of ECM proteins by endothelial progenitor cellscompared to two-dimensional cultures. These ECM proteins include forexample, fibronectin, laminin, and/or collagen IV. The polymermicrofibers also induce increased deposition of ECM proteins byperivascular cells seeded on the microfiber. The perivascular cellspreferably are vascular smooth muscle cells and/or pericytes. Theperivascular cells deposit increased ECM proteins including for example,collagens type I, III and IV, fibronectin, laminin or elastin. Inembodiments, the increased ECM proteins deposited include fibronectin,laminin, elastin, and collagens type I, III, and IV.

Another embodiment includes a luminal multicellular microvascularstructure. The luminal multicellular microvascular structure is formedfrom an electrostretched polymer microfiber seeded with endothelialprogenitor cells and a second cell type, for example perivascular(mural) cells, wherein the polymer microfiber is degraded afterattachment of the cells and deposition of the extracellular matrix. Thepolymer microfiber may be degraded with an enzyme, such as plasmin.Preferably, the endothelial progenitor cell is an endothelial colonyforming cell, and the perivascular cell is a vascular smooth muscle cellor pericyte. In another embodiment, the luminal multicellularmicrovascular structure is hollow.

In the embodiments of the invention, the polymer is selected fromalginate, fibrin (fibrinogen), gelatin, hyaluronic acid, or combinationsthereof. In particular embodiments, the polymer is fibrin. In theembodiments, endothelial progenitor cells are endothelial colony formingcells, and the perivascular cells are vascular smooth muscle cells orpericytes. Endothelial colony forming cells deposit ECM proteins such asfibronectin, laminin, or collagen type IV. Vascular smooth muscle cellsdeposit ECM proteins such as collagen types I, III, IV, laminin, elastinand fibronectin. Pericytes deposit ECM proteins such as collagen typesI, III, IV, laminin, and fibronectin.

Embodiments of the invention include a method of sequentiallycontrolling microvascular vessel formation comprising the steps ofpreparing a tubular polymeric scaffold, for example a polymeric hydrogelmicrofiber comprising electrostretched polymer microfibers; seeding themicrofiber with endothelial progenitor cells; co-culturing theendothelial cell-seeded microfiber with a second cell type such asperivascular (mural) cells, and varying the growth factors used for eachstep. The endothelial cells deposit extracellular matrix that producesextracellular proteins that encircle the microfiber, and are orientedperpendicular to the cell orientation, along the fiber's circumference,wherein formation of microvasculature vessel is sequentially controlled.In a preferred embodiment, the polymer is alginate, fibrin (fibrinogen),gelatin, hyaluronic acid, or combinations thereof. In a particularembodiment, the polymer is fibrin. In a preferred embodiment, theendothelial progenitor cells are endothelial colony forming cells andthe perivascular cells are vascular smooth muscle cells or pericytes. Inanother embodiment, the method further comprises degrading themicrofiber after seeding and culture of cells and deposition ofextracellular matrix. Degrading the microfiber may be with an enzyme,such as plasmin.

Another embodiment includes a system for sequentially controllingmicrovascular vessel formation comprising a tubular polymeric microfibercomprising electrostretched polymer microfibers for forming a matrix forthe culture of cells that form the vasculature; endothelial progenitorcells seeded on the polymer microfiber for forming a vascular lining;and perivascular (mural) cells co-cultured with the endothelialcell-seeded microfiber for forming a mural cell layer, and withendothelial progenitor cells depositing extracellular matrix proteinsthat encircle the microfiber and are oriented perpendicular to the cellorientation along the fiber circumference, wherein a microvascularstructure is formed. In an embodiment, the polymer is alginate, fibrin(fibrinogen), gelatin, hyaluronic acid, or combinations thereof. In aparticular embodiment, the polymer is fibrin. In a preferred embodiment,the endothelial progenitor cells are endothelial colony forming cellsand the perivascular cells are vascular smooth muscle cells orpericytes.

One embodiment of the present invention is a microfiber having alongitudinally aligned nanotopography comprising biodegradable,electrostretched hydrogel polymer nanofibers with internal polymeralignment. The microfibers of the present invention are preferablybiodegradable. A biodegradable microfiber of the present invention willnot include chemicals or compounds that inhibit the biodegradable natureof the microfiber. Examples of chemicals or compounds that inhibit thebiodegradable nature of microfibers include ceramics. Nanofibers of thepresent invention may be substantially free of such ceramics includingzirconia, silica (SiO₂), quartz, sapphire, diamond, Y₂O₃, Al₂O₃, CaO,MgO, TiO₂, P₂O₅, CaF₂, B₂O₃, and Na₂O as examples. Microfiber maycomprise nanofibers that are parallel to each other that are typicallyin the form of a solid bundle or a sheet. A microfiber comprising asolid bundle of nanofibers has a diameter in the range of 0.1 to 100 nm.A sheet of nanofibers maybe shaped to form a circumference so that amicrofiber of the present invention is hollow or comprises a conduit inthe range of 20 micrometers to 20 mm. Microfibers of the presentinvention may have a diameter in the range from 100 μm to about 500 μmbased on the outer circumference of the microfiber. The hydrogel polymernanofibers of the present may have a water content of greater than 90%,greater than 95%, or greater than 98% of the nanofiber. The nanofibersof the present invention comprise one or more polymers selected from thegroup consisting of alginate, fibrin (fibrinogen), gelatin, hyaluronicacid, and a combination thereof.

Another embodiment of the present invention are microfibers furthercomprising endothelial progenitor cells including endothelial colonyforming cells, as an example, seeded on the polymer microfiber. Theendothelial progenitor cells are aligned longitudinally to the polymermicrofiber and endothelial colony forming cells deposit extracellularmatrix proteins on the microfibers. The extracellular matrix proteinsthat are deposited are circumferentially organized, wrapping around themicrofiber. Examples of extracellular matrix proteins that are depositedinclude laminin, collagen IV, and fibronectin, as examples. The collagenIV, laminin, and fibronectin are deposited in higher quantities on themicrofiber than on 2D cultures. Other cells may adhere to themicrofibers of the present invention. For example, perivascular cellsmay be seeded, or adhere, on the microfiber. An example of suitableperivascular cells are pericytes. Pericytes deposit extracellular matrixproteins, and the extracellular matrix proteins are longitudinallyorganized along the microfiber. Another example of perivascular cellsare vascular smooth muscle cells. The vascular smooth muscle cellsdeposit extracellular matrix proteins, and these extracellular matrixproteins are longitudinally, randomly, or circumferentially organizedalong the microfiber. A second cell type may be seeded or adhere on themicrofiber. An example of a second cell type is a mural cell and themural cell is vascular smooth muscle cell or a pericyte. Microfibers ofthe present invention may comprise vascular smooth muscle cells orpericytes that is randomly oriented, or is longitudinally oriented withrespect to the microfiber. The vascular smooth muscle cell depositscollagen type I and elastin, and the pericyte deposits collagen type IV.These extracellular matrix proteins are induced to circumferentiallyorganize and wrap around the microfiber by the longitudinally alignednanotopography of the microfiber. Microfiber of the present inventionmay be substantially free of a chemical that promotes cell alignmentinclude hydrophilic agents such as dextran, polyvinyl alcohol,polyethylene glycol, polyoxyethylene, gelatin, pullan, heparin, hirudin,ticlopidine, and chlopidogrel; and hydrophobic agents such aspolyactide, polyactic acid, polyglycolide, polyglycolic acid,polyactide-polyglycolide, polyglycolide, polyglycolic acid,polyactide-polyglycolide, polycaprolactone, and polyamino acid, asexamples. The longitudinally aligned nanotopography of a microfiber ofthe present invention induces cell alignment on the microfiber. Achemical agent that promotes cell alignment in not needed on amicrofiber of the present invention in order to produce cell alignment.

Another embodiment of the present invention is a biodegradable,electrostretched hydrogel polymer nanofiber with internal polymeralignment. Nanofibers of the present invention also have alongitudinally aligned nanotopography and similar cell alignmentcharacteristics as the microfibers described above. Cells and proteinsalign on the nanofibers because of their longitudinally alignednanotopography Nanofibers of the present invention may be substantiallyfree of a chemical that promotes cell alignment include hydrophilicagents such as dextran, polyvinyl alcohol, polyethylene glycol,polyoxyethylene, gelatin, pullan, heparin, hirudin, ticlopidine, andchlopidogrel; and hydrophobic agents such as polyactide, polyactic acid,polyglycolide, polyglycolic acid, polyactide-polyglycolide,polyglycolide, polyglycolic acid, polyactide-polyglycolide,polycaprolactone, and polyamino acid, as examples. The nanofibers of thepresent invention are biodegradable and are substantially free of aceramics as describe above. The hydrogel polymer nanofibers has a watercontent of greater than about 90%, greater than 95%, or greater than98%.

BRIEF DESCRIPTION OF THE FIGURES

Having thus described the presently disclosed subject matter in generalterms, reference will now be made to the accompanying Figures, which arenot necessarily drawn to scale, and wherein:

FIGS. 1A-1M show the electrostretching setup and features of thepresently disclosed hydrogel microfibers: (a) illustration of arepresentative spinning setup for electrostretching; (b) effect ofalginate solution feeding rate on the diameter of the hydrogelmicrofibers. Alginate hydrogel microfibers with an average of 17-116 μmwere prepared with a solution containing 2% sodium alginate and 0.2% PEGfed at a flow rate ranging from 0.7-7.0 ml/h. Bars represent mean±s.d.(n=3); (c-f) demonstrate that various crosslinking mechanisms have beenemployed to crosslink alginate, gelatin, fibrin, and hyaluronic acidhydrogel microfibers. The crosslinking of the fibers was initiated witha fast calcium gelation of alginate, followed by additional crosslinkingof the second component polymer with UV-initiated, enzymatic, or theMichael addition reaction for methylated gelatin, fibrin and hyaluronicacid, respectively; (g) using this method, hydrogel microfibers of anydesired length can be prepared; (h) when dispersed in water, alginatehydrogel fibers formed a loose network of hydrogel fibers. Trypan bluewas used to stain the fibers and enhance observation; (i) a 10-g metalweight was lifted with an alginate hydrogel microfiber bundle; (j) amicro-knot was made with two alginate hydrogel microfibers; (f) under across polarized light microscope, light extinction was observed at thecrossover point of two hydrogel microfiber bundles, indicating uniformalignment in both fibers; (l-m) beyond microfiber bundles, thesehydrogel microfibers also can be fabricated into other forms likefibrous films (l) and self-supporting hydrogel tubes (m). Scale barsrepresent 100 μm in (c-f), 1 cm in (g-i) and (l-m), 50 μm in (j), and 1mm (k);

FIGS. 2A-2I show SEM micrographs of hydrogel fibers prepared with simpleextrusion and electrostretching. (a-c) Fibrin (a), gelatin (b) and HA(c) hydrogels prepared by simple extrusion or mixing consist of randomlyoriented nanofiber network. (d-f) Electrostretched fibrin (d), gelatin(e) and HA (f) hydrogel fibers showing preferential alignment. Arrowsindicate the orientation of the microfiber longitudinal axis. (g-i)Fibrin (g), gelatin (h) and HA (i) hydrogel fibers following stretchingand dehydration in air forming fiber bundles. Both fibrin and gelatinfibers preserved surface texture and grooves. Samples in (a-f) wereprepared by the critical point drying technique; and samples in (g-i)were stretched and dried in air. Scale bars represent 1 μm in (a-b) and(d-e), 2 μm in (c) and (f), 20 μm in (g-h), and 40 μm in (i);

FIGS. 3A-3G show X-ray scattering diffraction patterns and tensilemoduli of hydrogel fibers in dry and wet states. (a-b) Small angle X-rayscattering (SAXS) patterns of the dry (a) and wet (b) calcium alginatehydrogel fibers confirming an alignment axis along the microfiberorientation indicated by the arrows. (c) SAXS pattern of alginatehydrogel prepared by hand extrusion suggesting an isotropic structure.(d) Wide angle x-ray scattering pattern of the dry alginate microfibersconfirming the polymer chain alignment along the fiber axis as indicatedby the arrow. (e-g) Tensile moduli of alginate (AG), fibrin (FN),gelatin (GT) and hyaluronic acid (HA) hydrogel fibers in dry (e), wet(f) and re-hydrated form (g). Bars represent mean±s.d. (n=3);

FIGS. 4A-4F show small angle x-ray scattering (SAXS) patterns for fibrinand gelatin hydrogel microfibers: (a-b) SAXS patterns of dry (a) and wet(b) fibrin hydrogel fibers prepared by electrostretching; (c) SAXSpattern for fibrin hydrogel samples prepared by simple extrusion; (d-e)SAXS patterns of dry (d) and wet (e) gelatin hydrogel fibers prepared byelectrostretching; and (f) SAXS pattern for gelatin hydrogel samplesprepared by simple extrusion;

FIG. 5 shows an illustration of polymer alignment as a result ofelectrical and mechanical stretching. An aqueous solution ofbiopolymer(s) with or without cells is spun under electrical andmechanical stretching forces. Polymer chain alignment induced duringthis process is then quickly fixed with stabilizing solution in thecollection bath. Bicomponent or multicomponent hydrogel fibers can bespun similarly by mixing different polymers in the spinning solution,also referred to herein as the starting solution. Additionalcrosslinking can be performed via enzyme, UV-initiated crosslinking, orcell compatible chemical reactions (e.g. the Michael addition reaction).Cell encapsulation can be achieved by incorporating cells in thespinning solution, forming “cellular strings”; and

FIG. 6 shows tensile moduli of wet alginate hydrogel fibers prepared atdifferent collection plate rotation speeds. All fiber samples werecrosslinked in 25 mM CaCl₂) solution for 4 minutes prior to measurement.Bars represent mean±s.d. (n=3).

FIG. 7A-7E ECFCs attached and aligned on fibrin hydrogel microfibers.(A) Schematic of experimental procedure including electrostretching,ECFC seeding, and tumbling. Drawing not to scale. Scanning electronmicroscopy (SEM) of critical point-dried fibrin microfiber showingaligned topography on the microfiber surface. Scale bar is 10 μm. (B-D)Confocal Z-stack image reconstructions of ECFCs seeded on fibrinhydrogel microfibers horizontally aligned after 5 days in culture;F-actin (phalloidin staining) is shown in green, EC-specific markers(VECad, CD31, or vWF) in red, and nuclei in blue. Yellow arrows indicatethe direction of stretching and nanotopography on microfiber surface.Scale bars are 50 μm. n≥2 per stain with quadruplicates. (E) ConfocalZ-stack image reconstructions of ECFCs seeded on fibrin microfibersafter one day in culture; F-actin (phalloidin staining) are shown ingreen, CD31 in red, and nuclei are counterstained in blue. Scale barsare 100 μm.

FIG. 8A-8G ECFCs deposit ECM circumferentially on fibrin hydrogelmicrofibers. Confocal stack image reconstructions of ECFCs on fibrinmicrofibers after (A) 1 and (B) 5 days in culture. Scale bars are 200μm. (C) High magnification confocal images of laminin, fibronectin andcollagen IV wrapping around the fibrin microfibers. (D) TEM images ofcross-sectional slices of a cell-fibrin microfiber construct after 5days in culture (i-ii) with cells and (iii) without cells. (ii) is ahigher magnification image for the boxed area in (i). F=Fibrin; E=ECM;C=Cells. (E) Cross-sectional projections of confocal Z-stack images ofECFCs on fibrin microfibers after 5 days. F-actin (phalloidin) is shownin green, ECM proteins (collagen IV, laminin, fibronectin) in red ormagenta, and nuclei in blue. Yellow arrows indicate the direction ofnanotopography on microfiber surface. a-n≥2, b-e n≥5 per stain withquadruplicates. (F) and (G) High magnification confocal Z-stack imagereconstructions of ECFCs-seeded fibrin microfibers after 5 days inculture showing (F) wrapping ribbon-like organization of Collagen IV,laminin and fibronectin (in red; Scale bars are 50 μm) and (G)horizontal orientation of ECFCs with circumferential organization of thedeposited Collagen IV (red). Yellow arrow indicates the direction ofnanotopography. Scale bars are 100 μm.

FIG. 9A-G Nanotopography and geometry differently effect ECMorganization. Confocal Z-stack image reconstructions of (A) ECFCs on 2Dfibrin sheets after 5 days in culture. Yellow arrows indicate thedirection of nanotopography. Scale bars are 50 μm. n=2 with duplicates.(B) ECFCs on PES 3D fibrin-coated fibers with random non-alignedtopography after 5 days in culture. Scale bars are 100 n≥4 withquadruplicates. Actin filaments (phalloidin) are shown in green, ECMproteins (collagen IV, laminin, fibronectin) in red, and nuclei in blue.Box-and-whisker plots showing ECFCs (C) and ECM (D) angle of orientationon PES and fibrin hydrogel microfibers after 5 days in culture. (E)Standard deviation of ECM angle of orientation. Error bars represent5-95% confidence intervals. Significance levels in the mean representedby ***p<0.001. n≥2 with quadruplicates. (F) SEM of critical-point driedfibrin fiber sheets showing aligned topography on the surface. Scale baris 2 μm. (G) SEM of critical-point dried PES microfibers (i) coated withfibrin showing random topography and (ii) uncoated showing smoothtopography. Scale bars are 10 μm.

FIG. 10A-N Disrupting actin and microtubule organization does not affectECM organization. Confocal Z-stack image reconstructions of ECFCs seededon fibrin microfibers for 24 hrs followed by treatment with (A)cytochalasin D or (B) nocodazole for 24 hrs and 48 hrs in culture. (C)Low (left) and high (right) magnification of ECFCs seeded on fibrinmicrofibers for 72 hrs without drug treatment, serving as control.F-actin (phalloidin staining) is shown in green, microtubules(α-tubulin) in red, ECM proteins (collagen IV or fibronectin) in red ormagenta, and nuclei in blue. Yellow arrows indicate the direction ofnanotopography. Scale bars are 100 μm except of high magnification in(C) that is 50 μm. (D) Box-and-whisker plots and (E) standard deviationof ECM angle of orientation. Error bars represent 5-95% confidenceintervals. n=2 with quadruplicates. (F)-(H) Confocal Z-stack imagereconstructions of ECFCs seeded on (F) fibrin microfibers or (G) onPetri dish for 24 h followed by treatment with cytochalasin D for 48 hin culture. (H) ECFCs seeded on fibrin microfibers and treatedimmediately with cytochalasin D for 72 hrs of culture. F-Actin filaments(phalloidin) are shown in green, collagen IV in red, fibronectin orlaminin in magenta, and nuclei in blue. Yellow arrows indicate thedirection of nanotopography on fibrin microfibers. Scale bars are 50 μmin (F)-(G) and 100 μm in (H). (I)-(K) Confocal Z-stackhigh-magnification image reconstructions of ECFCs seeded on (I) fibrinmicrofibers or (J) Petri-dishes for 24 h followed by treatment withnocodazole for 48 h of culture. (K) ECFCs seeded on fibrin microfibersand treated immediately with nocodazole for 72 hrs of culture. F-Actinfilaments (phalloidin) in green, microtubules (α-tubulin) in red,Collagen IV in magenta, and nuclei in blue. Yellow arrows indicate thedirection of nanotopography on fibrin microfibers. Scale bars are 50 μmin (I) (left), (J) (right) and (K) (right); 20 μm in (I) (right); and100 μm in (K) (left). (L)-(N) Confocal Z-stack image reconstructions atdifferent magnifications of ECFCs-seeded fibrin microfibers after 3 daysin culture showing non-confluent ECFCs with circumferential organizationof the deposited Collagen IV (red). Yellow arrow indicates the directionof nanotopography. Scale bars are (L) 200 μm, (M) 100 μm, (N) 50 μm.

FIG. 11A-F Microfiber curvature influences ECM organization. (A)Confocal Z-stack image reconstructions of Collagen IV deposition onfibrin microfibers with different diameters. Scale bars are 200 μm (B)Scatter plot and (C) standard deviation of ECM angle of orientation onmicrofibers with different diameters. Error bars represent 5-95%confidence intervals. Significance levels in the distributionrepresented by ***p<0.001. n=2 with quadruplicates. (D)-(F) Highmagnification confocal Z-stack image reconstructions of ECFCs-seededfibrin microfibers with different diameter after 5 days in cultureshowing Collagen IV in red. Scale bars are 50 μm. Yellow arrow indicatesthe direction of nanotopography.

FIG. 12A-G Co-cultured vSMCs deposit new ECM. Confocal Z-stack imagereconstructions of fibrin microfibers seeded with ECFCs followed by (A)co-culture of vSMCs for 2 days. n=2 with quadruplicates. Scale bars are200 μm. Co-cultured vSMCs for 3 days showing (B) wrapping and (C)aligned arrangement. Scale bars are 100 μm. (D) Collagen Type Ideposited by co-cultured vSMCs after 3 days in co-culture. Scale barsare 100 μm. (E) Cross-sectional projection of confocal Z-stack images ofvSMCs after 5 days in co-culture. Arrowheads indicate SM22⁻ cells. Scalebars are 50 μm. (B)-(E) n≥3 with quadruplicates. (F) Confocal Z-stackimage reconstruction and (G) cross-sectional projection of co-culturedvSMCs after 5 days in co-culture. n=2 with quadruplicates. Scale barsare 50 SM22 is shown in green, CD31 in red, collagen I and elastin inred, and nuclei in blue.

FIG. 13A-E Co-cultured Pericytes proliferate and deposit new ECM.Confocal Z-stack image reconstructions of fibrin microfibers seeded withECFCs and cultured for 5 days followed by co-culture of pericytes for 10days in ECFC media supplemented with 30 mM aminocaproic acid. (A)Pericytes express SM22 (red) and are located above ECFCs expressinglectin (green). Co-cultured pericytes showing (B) wrapping and (C)aligned arrangement. SM22 shown in green. (D) Projection and (E)cross-sectional projection of confocal Z-stack images of Collagen IV(green) deposited by co-cultured pericytes marked with smooth muscleactin (SMA) (red). Nuclei shown in blue. Arrowheads indicate ECFCs whichare SM22⁻ and SMA⁻ (SMA negative) cells. Scale bars are 100 μm in (A)and (C), 50 μm in (B), (D), and (E).

FIG. 14A-C Deposition of Collagen IV (Col IV), fibronectin (Fn), andlaminin (Lmn) by ECFCs in 3D vs. 2D. Confocal Z-stack image projectionsof ECM in (A) 3D fibrin microfibers and (B) 2D fibrin coated surfaces.Red: corresponding ECM; blue: nuclei, green: F-actin. Scale bars=100 μm.Insert scale bars=25 μm. (C) RT-PCR analysis of expression of ECM genesby ECFCs cultured on 2D vs. 3D. Error bars represent SEM. Significancelevels in the distribution represented by *p<0.05 and **p<0.01. n≥3.

FIG. 15 A-D Deposition of Col I, III, IV, Fn, and Lmn by pericytes in 3Dvs 2D. Confocal Z-stack image projections of ECM in (A) 3D fibrinmicrofibers and (B) 2D fibrin coated surfaces. Red: corresponding ECM;blue: nuclei, green: F-actin. Scale bars=100 μm. Insert scale bars=25μm. Arrows, arrowheads, and double-headed arrows point to randomlydeposited, non-polymerized, and aligned ECM proteins, respectively. (C)RT-PCR analysis of expression of ECM genes by vSMCs cultured on 2D vs3D. (D) Orthogonal view of pericytes grown on microfibers. Arrowheadspoint to cells underneath outer cell layer, arrow points toextracellular deposited Lmn. Scale bar=20 μm. Error bars represent SEM.Significance levels in the distribution represented by *p<0.05 and**p<0.01. n≥3.

FIG. 16A-D Deposition of Col I, III, IV, Eln, Fn, and Lmn by vSMCs in 3Dvs 2D. Confocal Z-stack image reconstructions of ECM in (A) 3D fibrinfibers and (B) 2D fibrin coated surfaces. Red: corresponding ECM; blue:nuclei, green: F-actin. Scale bars=100 μm. Insert scale bars=25 μm.Arrows, arrowheads, and double-headed arrows point to randomlydeposited, intracellular, and aligned ECM proteins, respectively. (C)RT-PCR analysis of expression of ECM genes by pericytes cultured on 2Dvs 3D. ND=not determined. (D) Orthogonal view of pericytes grown onmicrofibers. Arrowheads point to cells underneath outer cell layer,arrow points to extracellular deposited Lmn. Scale bar=20 μm. Error barsrepresent SEM. n≥2.

FIG. 17A-C Fiber degradation and viability of cells and ECM afterplasmin treatment. (A) Light microscopy images of plasmin degradingfibrin microfibers in a concentration dependent manner. Scale bars=100μm. (B) Immunofluorescence images of viability assays of ECFCs after 12and 24 hr treatments with plasmin. Red: dead cells; green: live cellsScale bars=500 μm. (C) Confocal Z-stack image projections of fibrinfibers with ECFCs for 5 days and treated with plasmin for (I) 12 and(II) 24 hrs. (I) Col IV (red) Fn (green) Scale bars=200 μm, insert scalebars=100 μm. (II) Lmn (green) Left panel scale bar=200 μm, right panelscale bar=50 μm. Insert: Cross-sectional image, scale bar=50 μm. n≥2.

FIG. 18A-D ECFC and perivascular cell co-cultures on fibrin microfibers.Confocal Z-stack image projections of ECFCs grown for 5 days on fibrinmicrofibers followed by 5 days more of (A) pericyte co-culture. Red: (I)VEcad (II) SM22 (III) Col III; blue: nuclei, green: F-actin. (B) vSMCco-culture. Red: (I) VEcad (II) SM22 (III) Eln; blue: nuclei, green:F-actin. Confocal Z-stack image 3D reconstructions of structurescultures for 5 days with ECFC followed by 5 more days with perivascularcells and then treated for 12 hrs with 0.25 CU/mL plasmin. (C) Pericyteco-culture. Red: (I) and (III) VEcad, (II) and (IV) Col III; magenta:(I) and (III) SM22; blue: nuclei; green: (I) and (III) F-actin, (II) and(IV) Col IV. (D) vSMC co-culture. Red: (I) and (III) VEcad, (II) and(IV) Eln; magenta: (I) and (III) SM22, (II) and (IV) Fn; blue: nuclei;green: F-actin. Scale bars=50 μm. n≥2.

DETAILED DESCRIPTION

The presently disclosed subject matter now will be described more fullyhereinafter with reference to the accompanying Figures, in which some,but not all embodiments of the inventions are shown. Like numbers referto like elements throughout. The presently disclosed subject matter maybe embodied in many different forms and should not be construed aslimited to the embodiments set forth herein; rather, these embodimentsare provided so that this disclosure will satisfy applicable legalrequirements. Indeed, many modifications and other embodiments of thepresently disclosed subject matter set forth herein will come to mind toone skilled in the art to which the presently disclosed subject matterpertains having the benefit of the teachings presented in the foregoingdescriptions and the associated Figures. Therefore, it is to beunderstood that the presently disclosed subject matter is not to belimited to the specific embodiments disclosed and that modifications andother embodiments are intended to be included within the scope of theappended claims.

The presently disclosed subject matter provides an approach thatcombines an electrical and mechanical stretching force to generatebiopolymer hydrogel microfibers having a high degree of chain alignmentwithin the hydrogel fiber. The presently disclosed methods can beapplied to a wide range of polymer hydrogel systems, such as alginate,fibrin, gelatin, collagen, hyaluronic acid, chitosan, and their blendsand are applicable to a wide range of biomedical applications.

The presently disclosed subject matter demonstrates that this internalpolymer chain alignment affords excellent mechanical properties to thesehydrogel fibers. The presently disclosed methods are highly versatilewith a high degree of control over fiber diameter and fiber constructs.Further, the presently disclosed fiber spinning process is conducted inaqueous solutions at room temperature and is thus amenable to cellencapsulation within the hydrogel fibers during spinning and gelation.

Further, the presently disclosed subject matter demonstrates that thealignment topography is a strong matrix cue to induce alignment of cellsthat are seeded either inside the hydrogels or on the hydrogel surface.Due to the excellent cellular responses and the versatility of materialschoice, these systems can be useful substrates to create “cellularwires” (e.g., nerve cables) or guide cell migration in wound healing orregeneration.

I. Methods of Preparing Electro-Mechanically Stretched Microfibers

Current electrospinning methods known in the art for producingmicrofibers rely on electrical force to stretch the fiber into a smallerdiameter. In such methods, dried fiber is collected after an acceleratedstretching process, during which the speed and amount of jet elongationinduced by an applied electrical field is not well controlled. The highstretching rate (e.g., 10⁵ to 10⁶ s⁻¹) at the end of such processes alsomakes known electrospinning techniques unsuitable for treating delicatesubstances, such as hydrogel, cells, or self-assembled molecules.Further, large fibers, e.g., fibers having a diameter ranging from tensto hundreds of microns, are not easy to make by electrospinning methodsknown in the art.

A certain level of molecular orientation, however, can be developed viaconventional electrospinning processes. Molecular level alignment inthese cases is a result of high strain rate commonly used inelectrospinning of polymers (e.g., 10⁵ to 10⁶ s⁻¹). According totheoretical models, a high degree of uniaxial orientation is expected ifthe product of strain rate and the conformational relaxation time λ isgreater than unity during uniaxial stretching of polymeric melts orsolutions. Generally, a high level crystalline structure along the fiberaxis is not observed in fibers formed by conventional electrospinningprocesses because of the rapid solidification of the fluid jet.

In contrast, the presently disclosed methods use a combination ofelectrical and mechanical force to induce stretching to replace theuncontrollable stretching in electrospinning. More particularly, thepresently disclosed methods use an electrical field to initiate andstretch a jet stream of polymer solution and a mechanical force exertedby a rotating collection bath comprising a stabilizing collectionsolution and a rotating collection plate to control the amount ofstretching. In this way, the overall stretching rate can be adjusted bychanging the speed of the rotating collection plate and by modifyingelectric field strength. Fibers produced this way also have a molecularlevel preferential alignment, which significantly improves theirmechanical properties.

Generally, in the presently disclosed methods, a high voltage electrodeis contacted with a starting solution to initiate a jet stream ofpolymer solution through, for example, a syringe needle. The jet streamof polymer solution is collected with a rotating collection platepositioned at a close distance to the tip of the syringe needle beforethe jet stream of polymer solution is accelerated by an electrical fieldinduced by the high voltage electrode. More particularly, as providedherein above, the electric field initializes the stretching of thepolymer stretch, which is due to an acceleration of the jet stream ofpolymer caused by the applied electrical field. The collection of thejet stream of the polymer solution occurs while the stream is still in alinear trajectory before the chaotic bending instability (whipping)regime that is standard to traditional electrospinning.

During collection, the rotating collection plate further stretches thejet stream of polymer solution so it travels the same distance as therotating collection plate. Because the mechanical stretching rate isdetermined by the speed of the rotating collection plate and solutionfeed rate, the presently disclosed methods can be regulated to processdelicate substances or to produce fibers of desired size range (10 to300 μm), both of which cannot be done with conventional electrospinningmethods known in the art.

In the presently disclosed methods, the jet stream of polymer solutionis collected in regions where the jet stream is initiated and before itis accelerated into a whipping jet. The overall electromechanical strainrate is estimated to be around 10 to 70 s⁻¹, which is several orders ofmagnitude lower than conventional electrospinning. The relative highmolecular weight of natural polymers used in this process, correspondingto a longer relaxation time λ, will be sufficient to facilitate polymerchain alignment under low strain rate, in absence of rapidsolidification.

Accordingly, the elongational flow induced by electrical and mechanicalstretching force also creates a unique alignment among the individualfibers in the string or film produced. Without wishing to be bound toany one particular theory, it is thought that the synergy of electricalfield and mechanical stretching helps to align the fibers. In additionto stretching induced alignment, the electrical field also contributesto the overall alignment of the fibers. The alignment can be fixed, forexample by crosslinking the fibers after they are collected. Suchalignment not only enhances the mechanical properties of the finalmatrix, but also brings new capacities for their utility. Polarizedoptical microscopy and scanning electron microscopy (SEM) have been usedto confirm these characteristic structures.

Fibers formed by the presently disclosed can be further elongated withmechanical force along the fiber axis. Application of such mechanicalforce or stress helps to enhance the alignment of the polymer chains andfix them in a longitudinal direction. Due to their large surface areas,such strings will bundle together if taken out the collection or fixingsolution. Applying a constant stretching force until the fibers are drycan lead to a fiber cross sectional area that decreases to approximately2% of its original cross sectional area. Again, without wishing to bebound to any one particular theory, it is thought that such adehydration process further improves the alignment inside the string.

Accordingly, in some embodiments, the presently disclosed subject matterprovides methods to incorporate anisotropic topography inside a hydrogelmatrix using a combination of electrical and mechanical stretching. Suchmicrofibers can be made of natural polymers including, but not limitedto, alginate, fibrinogen, gelatin, collagen, hyaluronic acid (HA),chitosan chondroitin sulfate, dextran sulfate, heparin, heparan sulfate,and the like, and functionalized derivatives thereof; synthetic polymersincluding, but not limited to, polyacrylic acid derivatives, polyvinylalcohol, and the like.

The presently disclosed methods are versatile and scalable and representthe first approach to enable control over hydrogel alignment topographyin polymer hydrogel fibers derived from a variety of biopolymers.Accordingly, the molecules inside microfibers made by the presentlydisclosed methods are preferentially aligned along the microfiber axis.The unique internal uniaxial alignment characteristic enhances themechanical properties of the hydrogel microfibers. Microfibers having asize ranging from a few microns to hundreds of microns have been madeusing the presently disclosed electro-mechanical stretching method. Thepresently disclosed methods can be used to produce various forms ofhydrogel matrices, such as a film, mesh, tube, a single string, or abundled yarn.

Further, mechanical strain usually generates dense materials through theexclusion of water during the stretching process. In contrast, thepresently disclosed methods retain the hydration ratio or water contentthroughout the spinning process. As a result, hydrogel fibers with awater content of more than 90%, and, in some embodiments, at high as99%, including 90%, 91%, 92%, 93%, 94%, 95%, 96%, 97%, 98%, and 99%water content.

In other embodiments, twisted yarn can be produced from the hydrogelstrings described hereinabove. To make a yarn, a multiround of the loopis taken out of the fixing solution and hung in the air with a weightapplied at the bottom of the string. The string will elongate under suchweight. The string can be elongated much more when the stretching isdone at low dehydration degree. This process helps to further align thefibers. If this is done after significant crosslinking, then thedeformation is limited. For example, a dried yarn of alginate is verystrong and has a Young's modulus of up to 10 GPa. Such dried strings canbe rehydrated after being placed back into water.

Accordingly, in some embodiments, the presently disclosed subject matterprovides a method for preparing a microfiber having a uniaxialalignment, the method comprising: (a) providing at least one startingsolution comprising one or more polymers; (b) applying an electricalpotential to the at least one starting solution sufficient to initiate ajet stream of polymer solution; and (c) mechanically stretching the jetstream of polymer solution during or after collecting the jet stream ofpolymer solution in a collection bath comprising a stabilizing solution,wherein the collection bath is positioned at a separation distance suchthat the jet stream of polymer solution is collected before it isaccelerated by an electrical field created by the applied electricalpotential. In other embodiments, the collection bath comprises arotating collection plate, wherein a rotation of the collection platemechanically stretches the jet stream of polymer solution as it iscollected thereon. In still other embodiments, the collection bathcomprises a stationary collection plate, wherein the jet stream ofpolymer solution is deposited on the stationary collection plate using aback and forth motion to mechanically stretch the jet stream of polymersolution as it is deposited on the stationary collection plate. In someembodiments, one or more polymers comprise a natural polymer. In otherembodiments, the natural polymer is selected from the group consistingof water soluble polysaccharides, proteins, and combinations or blendsthereof. In particular embodiments, the natural polymer is selected fromthe group consisting of one or more of alginate, fibrinogen, gelatin,collagen, hyaluronic acid (HA), chitosan, chondroitin sulfate, dextransulfate, heparin, heparan sulfate, functionalized derivatives thereof,and combinations or blends thereof.

In yet other embodiments, one or more polymers comprise a syntheticpolymer. In some embodiments, the synthetic polymer is selected from thegroup consisting of a polyester and a polyamide. In other embodiments,the polyester is selected from the group consisting of polylactic acidand poly(lactic-co-glycolic) acid. In particular embodiments, thesynthetic polymer is selected from the group consisting of apolyacrylate, a poly(vinyl alcohol), a poly(ethylene glycol),functionalized derivatives thereof, and combinations or blends thereof.

In some embodiments, the starting solution further comprises athickening agent capable of increasing a viscosity of the jet stream ofpolymer solution. In particular embodiments, the thickening agentcomprises polyethylene glycol (PEG).

In other embodiments, the method further comprises crosslinking themicrofiber. In particular embodiments, the crosslinking is selected fromthe group consisting of ionic crosslinking, ultraviolet crosslinking,enzymatic crosslinking, and a chemical crosslinking reaction. In someother embodiments, the method further comprises adding a crosslinkingagent to the starting solution comprising one or more polymers. In stillother embodiments, the method further comprises adding a crosslinkingagent to the jet stream of polymer solution after the jet stream ofpolymer solution is initiated by the applied electrical potential. Infurther embodiments, the method comprises adding a crosslinking agent tothe collection bath.

In particular embodiments, the electrical potential has a range fromabout 2 kV to about 6 kV.

In some embodiments, the stabilizing solution comprises a solvent inwhich the jet stream of polymer solution is insoluble and precipitatesin the stabilizing solution. In other embodiments, the jet stream ofpolymer solution comprises an aqueous solution and one or morewater-soluble polymers and the stabilizing solution comprises an organicsolvent.

In yet other embodiments, the method further comprises elongating themicrofiber by applying mechanical stress along the uniaxial alignmentthereof and drying the microfiber.

In some embodiments, the method further comprises combining multiplemicrofibers to form a fiber bundle.

In other embodiments, the at least one starting solution comprises ablend of two different polymers; or two starting solutions are provided,wherein each starting solution comprises a different polymer; and themicrofiber comprises a bicomponent fiber having a core and a sheath.

In some embodiments, the method further comprises adding one or morebioactive agents to at least one starting solution. In otherembodiments, the method further comprises depositing one or morebioactive agents on the microfiber after it is formed.

In still other embodiments, the method further comprises adding one ormore cells to at least one starting solution. In further embodiments,the method further comprises seeding the microfiber with one or morecells on a surface of the microfiber after it is formed. In yet otherembodiments, the presently disclosed subject matter provides amicrofiber formed by the presently disclosed methods. In someembodiments, the microfiber has a diameter ranging from about 5 micronsto about 300 microns.

In some embodiments, the microfiber comprises more than one polymer. Inother embodiments, the microfiber comprises a bicomponent fibercomprising a core and a sheath.

In further embodiments, the microfiber comprises a hydrogel. In stillfurther embodiments, the hydrogel has a water content of greater thanabout 90%. In other embodiments, the hydrogel has a water content ofgreater than about 95%. In still other embodiments, the hydrogel has awater content of greater than about 98%.

In some embodiments, the microfiber has internal molecular chainalignment.

In other embodiments, the microfiber further comprises one or morebioactive agents. In still other embodiments, the microfiber furthercomprises one or more cells.

In other embodiments, bioactive agents may be either post-loaded in themicrofibers or loaded in situ within the microfiber as a component ofthe starting polymer solution, wherein the bioactive agents may include,but are not limited to, therapeutic agents, nanoparticles, water solubleproteins, cells, and their like. In still other embodiments, thebioactive agents are used for localized, sustained release in vitro orin vivo.

II. Use of Electro-Mechanically Stretched Microfibers as ScaffoldMaterials for Cell or Tissue Growth

A highly endeavored topic in regenerative medicine is to createextracellular matrix (ECM) analogs for providing mechanical supports andbiochemical cues to cells. For cases, such as tendons, nerves andcorneal stroma and intervertebral-disc regeneration, a matrix that canguide cellular alignment and growth direction is essential for optimalresults.

Many new scaffold materials have been developed for such purposes. Forexample, Zhang et al. have developed self-assembly peptide hydrogelstrings that can align encapsulated cells and others also have usedelectrospun nanofibers to guide cellular growth direction. Aligned ECMmade from natural polymer fibers, which in many cases possessunprecedented biological performance, are limited however tomicrofluidic alignment, cyclic mechanical stretching, and the like.

The facile, organic solvent-free processing conditions of the presentlydisclosed methods are amenable to the incorporation of live cells and/orgrowth factor proteins within the hydrogel fiber or on the fiber surfaceand effectively induce cellular alignment and provide cellular growthguidance.

Accordingly, in some embodiments, the presently disclosed microfiberhaving a uniaxial alignment can be used as a template for growing andguiding cells. The presently disclosed microfibers also can incorporateencapsulated cells and/or growth factor proteins. Such microfibers canbe used for making a cellular guide, a nerve guide for neuronalregeneration, a template for growing micro-blood vessels, surgicalsutures, wound dressing, or tissue scaffolds for tissue engineering.

In further embodiments, a co-axial spin of a hydrogel core/sheathstructure with cellular content in the core also can be produced.

In a representative example, the electro-mechanically stretched hydrogelstring can be used to direct the orientation of cells trapped inside.After dispersing mammalian cells in alginate/fibrinogen mixturesolution, the hydrogel strings can be stretched and collected in 50 mMCaCl₂) and 5-20 units/mL thromin solution to form hydrogel fibers withencapsulated cells. The facile spinning condition will ensure that theencapsulated cells remain viable during the process of fiber formationand culture.

In other embodiments, the same cells can be cultured on the surface, aswell. It is obvious to those skilled in the art that these cells canalign with the axis of the fiber.

Although specific terms are employed herein, they are used in a genericand descriptive sense only and not for purposes of limitation. Unlessotherwise defined, all technical and scientific terms used herein havethe same meaning as commonly understood by one of ordinary skill in theart to which this presently described subject matter belongs.

Following long-standing patent law convention, the terms “a,” “an,” and“the” refer to “one or more” when used in this application, includingthe claims. Thus, for example, reference to “a subject” includes aplurality of subjects, unless the context clearly is to the contrary(e.g., a plurality of subjects), and so forth.

Throughout this specification and the claims, the terms “comprise,”“comprises,” and “comprising” are used in a non-exclusive sense, exceptwhere the context requires otherwise. Likewise, the term “include” andits grammatical variants are intended to be non-limiting, such thatrecitation of items in a list is not to the exclusion of other likeitems that can be substituted or added to the listed items.

For the purposes of this specification and appended claims, unlessotherwise indicated, all numbers expressing amounts, sizes, dimensions,proportions, shapes, formulations, parameters, percentages, parameters,quantities, characteristics, and other numerical values used in thespecification and claims, are to be understood as being modified in allinstances by the term “about” even though the term “about” may notexpressly appear with the value, amount or range. Accordingly, unlessindicated to the contrary, the numerical parameters set forth in thefollowing specification and attached claims are not and need not beexact, but may be approximate and/or larger or smaller as desired,reflecting tolerances, conversion factors, rounding off, measurementerror and the like, and other factors known to those of skill in the artdepending on the desired properties sought to be obtained by thepresently disclosed subject matter. For example, the term “about,” whenreferring to a value can be meant to encompass variations of, in someembodiments, ±100% in some embodiments ±50%, in some embodiments ±20%,in some embodiments ±10%, in some embodiments ±5%, in some embodiments±1%, in some embodiments ±0.5%, and in some embodiments ±0.1% from thespecified amount, as such variations are appropriate to perform thedisclosed methods or employ the disclosed compositions.

Further, the term “about” when used in connection with one or morenumbers or numerical ranges, should be understood to refer to all suchnumbers, including all numbers in a range and modifies that range byextending the boundaries above and below the numerical values set forth.The recitation of numerical ranges by endpoints includes all numbers,e.g., whole integers, including fractions thereof, subsumed within thatrange (for example, the recitation of 1 to 5 includes 1, 2, 3, 4, and 5,as well as fractions thereof, e.g., 1.5, 2.25, 3.75, 4.1, and the like)and any range within that range.

III. In Vitro Cellular and Extracellular Matrix (ECM)

A novel in vitro model and system that recapitulates key aspects in thecellular and extracellular matrix (ECM) organization of themicrovasculature is established in accordance with the invention. Thismodel and system guide the formation of organized microvascularstructures, induction of endothelial cell alignment and elongation, anddemonstrates circumferential deposition of ECM proteins by endothelialprogenitor cells, for example endothelial colony forming cells. Themodel reveals the role of vessel diameter on ECM organization duringhuman microvascular growth. The model supports a step-wise vascularformation process via introduction of perivascular cells and differentgrowth factors at varying time points, a current challenge inmicrovascular tissue engineering. A multicellular microvascularstructure with an organized endothelium and multicellular perivasculartunica media is also disclosed.

Most current approaches for the in vitro study of the microvasculaturein a 3D setting use hydrogels and scaffolds embedded with vascularcells, which are subject to spontaneous capillary bed formation(Hielscher A C, et al., Am J Physiology—Cell Physiology, 2012; 302:C1243-C1256; Soucy P A and Romer L H, Matrix Biology, 2009; 28: 273-283;Hanjaya-Putra D, et al., Blood, 2011; 118: 804-815, Moon J J, et al.,Biomaterials, 2010; 31: 3840-3847; Pham Q P, et al., Tissue Eng, 2006;12: 1197-1211; Benjamin L E, et al., Development, 1998; 125: 1591-1598;Gerhardt H, Betsholtz C, Cell and Tissue Research, 2003; 314: 15-23;Stratman A N, et al, Blood, 2009; 114: 5091-5101; Wang Z Z, et al., NatBiotech, 2007; 25: 317-318; Koike N, et al., Nature, 2004; 428: 138-139;Melero-Martin J M, et al., Circulation Research, 2008; 103: 194-202;Levenberg S, et al., Nature Biotechnology, 2005; 23: 879-884; HielscherA C, Gerecht S, Cancer Research, 2012; 72: 6089-6096; Soucy P A, et al.,Acta Biomaterialia, 2011; 7: 96-105; Hanjaya-Putra D, et al.,Biomaterials, 2012; 33: 6123-6131; Leslie-Barbick J E, et al.,Biomaterials, 2011; 32: 5782-5789), or use micropatterned hydrogels togenerate organized microvasculature structures (Baranski J D, et al.,Proc Natl Acad Sci USA, 2013; Miller J S, et al., Nat Mater, 2012; 11:768-774; Zheng Y, et al., Proc Natl Acad Sci USA, 2012; 109: 9342-9347).While these approaches are instrumental for studying angiogenicprocesses, they provide only partial control over the topographical cuespresented to the cells by the ECM and possess limited opportunities tocreate and investigate multi-cellular vascular structures with properECM organization.

Previous studies in the field of angiogenesis, vasculogenesis, andvascular tissue engineering have either focused on studying theformation of capillaries and vessels with diameters below 100 μm(Hanjaya-Putra, D., et al., J Cellular and Molecular Medicine, 2010;14(10):2436-2447; Hanjaya-Putra, D., et al., Blood, 2011; 118:804-815;Davis, G. E., et al., Birth Defects Research Part C: Embryo Today:Reviews, 2007; 81(4):270-285; Montaño, I., et al., Tissue EngineeringPart A, 2009. 16(1): p. 269-282; Chen, X., et al., Tissue EngineeringPart A, 2008. 15(6): p. 1363-1371; Kobayashi, A., et al., Biochemicaland Biophysical Research Communications, 2007. 358(3): p. 692-697;Dickinson, L. E., et al., Soft Matter, 2010. 6(20): p. 5109-5119; Tsuda,Y., et al., Biomaterials, 2007. 28(33): p. 4939-4946; Baranski, J. D.,et al., PNAS, 2013. 110(19): p. 7586-7591), or have instead aimed todevelop large-diameter tissue engineered vessels, most over 3 mm indiameter (Vaz, C. M., et al., Acta Biomaterialia, 2005. 1(5): p.575-582; Kelm, J. M., et al., J Biotechnology, 2010. 148(1): p. 46-55;L'heureux, N., et al., FASEB Journal, 1998. 12(1): p. 47-56; Dahl, S.L., et al., Cell transplantation, 2003. 12(6): p. 659-666; Quint, C., etal., PNAS, 2011. 108(22): p. 9214-9219).

The invention overcomes challenges to developing physiologicallyrelevant microvascular structures with diameters between 100 μm and 1mm. It previously has been established that ECs can create vascularnetworks in vitro when cultured in 3D matrices such as hydrogels, yetthese networks result in capillary beds with relatively small lumendiameters (Hanjaya-Putra, D., et al., J Cellular and Molecular Medicine,2010; 14(10):2436-2447; Hanjaya-Putra, D., et al., Blood, 2011;118:804-815; Davis, G. E., et al., Birth Defects Research Part C: EmbryoToday: Reviews, 2007; 81(4):270-285; Montaño, I., et al., TissueEngineering Part A, 2009. 16(1): p. 269-282; Chen, X., et al., TissueEngineering Part A, 2008. 15(6): p. 1363-1371). Therefore, to createvessels with a larger diameter the present invention guides theformation of cylindrical vascular structures in the order of hundreds ofmicrons. Microfluidic channels have been used extensively to studymicrovascular development processes, yet most of these studies are donein channels with a square cross-section or in non-implantable devices(Verbridge, S. S., et al., J Biomedical Materials Research Part A, 2013.101(10):2948-2956; Abaci, H. E., et al., Sci. Rep., 2014:4).

Recent studies have developed 3D microfluidic channel arrays withrectangular (Zheng, Y., et al., PNAS, 2012; 109(24):9342-9347) orcircular cross-sections embedded in hydrogels (Miller, J. S., et al.,Nature Materials, 2012; 11(9):768-774; Wu, W., et al., AdvancedMaterials, 2011; 23(24):H178-H183; Kolesky, D. B., et al., AdvancedMaterials, 2014; 26(19):3124-3130; He, J., et al., Adv. HealthcareMaterials, 2013; 2(8): p. 1108-1113). However, these present limitedsuccess for constructing a multilayered structure with a continuousendothelium and a robust supporting multicellular mural cell layer. Toaddress the need for mural cell involvement, some of these studiesincorporated perivascular cells in the hydrogels encompassing themicrofluidic channels, and afterwards seeded ECs in the lumen (Zheng,Y., et al., PNAS, 2012; Miller, J. S., et al., Nature Materials, 2012).While this approach allows the study of cell recruitment, it imposes abarrier for full investment of perivascular cells, which have to migratethrough the hydrogels to find the developing microvessels. As such,perivascular cells in these systems were not demonstrated to form auniform multicellular layer on top of the endothelium. Furthermore,these systems impart a barrier for the detailed study of ECMorganization and EC-mural cell interactions due to chemical and physicallimitations presented the hydrogels.

The microfiber according to the invention provides control over thetopographical cues presented to cells by the ECM. The microfiber of theinvention also presents opportunities to create and investigatemulti-cellular vascular structures with proper ECM organization. Themicrofiber according to the invention has a nanotopography that induceslongitudinal adhesion and alignment of endothelial progenitor cells, forexample, endothelial colony-forming cells (ECFCs). The endothelialprogenitor cells, such as ECFCs, deposit circumferentially organizedECM. The ECM wraps around the microfibers of the invention, which isindependent of ECFCs' actin and microtubule organization. As establishedby the present invention, ECM encircling or wrapping around themicrofibers is dependent on the curvature of the microfiber. Accordingto the invention, microfibers with small diameters, for example lessthan about 500 μm, preferably about 100 μm to about 450, more preferablyabout 100 μm to about 370 μm, guide circumferential ECM deposition.Microfibers with larger diameters, 445 μm and higher, do not supportwrapping ECM as effectively. The invention provides a novel in vitrostructure and method for the sequential control of microvasculaturedevelopment and reveals the unprecedented role of the endothelium inorganized ECM deposition regulated by the microfiber curvature.

Unless defined otherwise, technical and scientific terms used hereinhave the same meaning as commonly understood by one of ordinary skill inthe art to which this invention belongs. As used herein, the terms“perivascular cells” and “mural cells” have the same meaning. The term“about” when referring to a measurable value such as an amount, atemporal duration, and the like, is meant to encompass variations of±20% or ±10%, more preferably ±5%, even more preferably ±1%, and stillmore preferably ±0.1% from the specified value, as such variations areappropriate to perform the disclosed methods.

The present invention can be used as a model of the microvasculaturethat recapitulates both cellular and ECM organization, towards theunderstanding of microvasculature development and utilization of themodel for regenerative medicine applications. Towards this,electrostretched microfibers designed to generate a micro-cylindricalmold with a line-grating nanotopography are used to enable bothendothelial layer organization and co-culture of supporting perivascular(mural) cells, such as vascular smooth muscle cells (vSMCs) orpericytes. Microfibers used have diameters of about 500 μm or less,preferably ranging from about 100 μm to about 450 μm, corresponding to apoorly studied range of vasculature in the body, namely venules andarterioles. Furthermore, in the existing models of microvasculature, thedeposition and organization of ECM proteins by endothelial cells has notbeen studied. Moreover, the full investment of mural cells on theendothelium of microvascular models has been challenging, due in part tothe use of endothelial-lined void spaces in most models (Miller J S, etal., Nat Mater, 2012; 11: 768-774; Zheng Y, et al., PNAS USA, 2012; 109:9342-9347), which introduces a cell migration barrier for mural cellinvestment. In contrast, the model of the invention allows not only highresolution studies of both cell and ECM organization; it allowsintroduction of mural cells after endothelial layer formation andenables full investment of these mural cells, recreating the media layerof microvasculature.

This new approach to create aligned hydrogel microfibers as describedherein uses an electrostretching process from various polymer materials.Unique characteristics of the electrostretched polymer fibers are theinternal and topographical alignment of the fibrous structure, generatedas a result of both electrical field and mechanical shear-inducedpolymer chain alignment. Furthermore, the microfiber diameter iscontrollable and uniform as a result of the bundling and processing ofthe individual fibers composing the microfibers.

The typical electrospinning process, such as disclosed in InternationalPublication WO2013/165975, incorporated by reference herein, consists ofa syringe pump with a syringe containing a polymer solution of one ormore polymers, a high voltage source, and a grounded collecting plate.The technique is based on inducing an electric charge on the polymersolution while applying an electric field between the syringe needle andthe grounded collecting plate. As the solution is dispensed from thesyringe, the electrostatic force opposes the surface tension of thepolymer solution producing a Taylor cone. Eventually the electrostaticforce overcomes the surface tension to produce a liquid jet stream. Asthe jet travels, the electric forces cause the stream to spin. At thesame time, the solvent evaporates from the solution and the polymerfibers fall on the collecting plate forming an ultrathin nanofiber. (SeeWO2013/165975).

The novel electrospinning/electrostretching technique of the inventionmodifies the typical electrospinning processes, and produces hydrogelmicrofibers with a uniaxial aligned topography using a combination ofelectrical and mechanical stretching. The typical electrospinningprocess was modified so the collecting plate was a grounded rotatingdisc containing an aqueous solution (See FIG. 7 ). The polymer jetemitted from the syringe is deposited as nanofibers that fall on top ofeach other in the rotating disc bath. At this point, the bundle ofaligned nanofibers is not cohesive. To make a cohesive microfiber, theelectrospun nanofibers are collected together parallel to each other andstretched mechanically by the rotating disc, then air-dried so thenanofiber bundle becomes the microfiber. This bundle is furtherstretched to obtain a cohesive uniaxial internal and topographicalalignment, which enhances the mechanical properties of the microfibers.Alternatively, the nanofibers can be collected together and partiallystretched on a modified plastic frame to make a flat 2D nanofiber sheet.

Various polymers may be used in the electrostretching technique of theinvention, such as natural polymers including alginate, fibrin(fibrinogen), gelatin, hyaluronic acid (HA), chitosan chondroitinsulfate, dextran sulfate, heparin, heparan sulfate, and functionalizedderivatives thereof, and synthetic polymers selected from a polyesterand a polyamide, such as polyacrylic acid derivatives and polyvinylalcohol, including polylactic acid, poly(lactic-co-glycolic) acid,polyacrylate, poly(vinyl alcohol), poly(ethylene glycol), as well ascombinations thereof, that produce hydrogel polymer fibers useful in theinvention. Nanofibers formed from the polymer may be crosslinked.Crosslinking may be achieved by any cross-linking method, includingionic crosslinking, ultraviolet crosslinking, enzymatic crosslinking,and chemical crosslinking reaction. For the electrostretching techniquepresented in the invention, the preferred polymers that can be used arealginate, gelatin, fibrin (fibrinogen), hyaluronic acid, andcombinations thereof, with fibrin being the most preferred polymer.

Fibrin gels have been used to study microvasculature assembly (DickinsonL E, et al., Soft Matter, 2010; 6: 5109-5119; Bayless, K J, and Davis, GE, Biochemical and Biophysical Research Communications, 2003; 312:903-913; Davis G E, and Bayless K J, Microcirculation, 2003; 10: 27-44;Bayless K J, et al., RGD-Dependent American Journal of Pathology, 2000;156: 1673-1683; Dickinson L E, et al., Lab Chip, 2012; 12: 4244-4248),vSMC responses (Ahmann K A, et al., Tissue Eng Part A, 2010; 16:3261-3270; Long J L, and Tranquillo R T, Matrix Biol, 2003; 22: 339-350)and multicellular organization (Lesman A, et al., Biomaterials, 2011;32: 7856-7869). Fibrin is used as a matrix material for the microfiberaccording to the invention, including use in preparing hydrogelmicrofibers as a template for the step-wise creation of microvasculatureof the invention.

The polymer microfiber of the invention has an aligned nanotopographythat guides alignment and elongation of endothelial progenitor cells,such as ECFCs. The invention provides a microfiber having cylindricalshape and tunable diameter of the fibers, which are novel features inhydrogels that allow detailed 3D view and analysis of microvasculaturedevelopment. The novel features also allow analysis of the effect ofcurvature of the fiber on cell processes. In an embodiment of theinvention, fibrin is used for the hydrogel microfiber, which makes thescaffold not only biocompatible, bio-adhesive, and pro-angiogenic, butalso easily degradable, such as through plasmin fibrinolysis. Thedevelopment of delimited microvascular structures in small size range,for example less than about 500 μm, preferably about 100 μm to 450 μm,with demonstrated detailed cell and ECM organization has not beenpreviously achieved.

According to the invention, aligned polymer microfiber (nanofiberbundles) used as a cylindrical platform control the organized adhesionof endothelial progenitor cells, such as ECFCs. In an embodiment of theinvention, endothelial progenitor cells like ECFCs are cultured onelectrospun fibrin microfibers that have a diameter of less than about500 μm, preferably 100 μm to about 450 μm. The cells attach throughoutthe microfibers. ECFCs form a continuous monolayer over the entiremicrofiber, with a distinctive elongated and mature morphology. Thepolymer microfibers offer an innovative approach in which ECFCs areseeded on the surface of an electrostretched microfiber, as opposed toseeded in the body of a nanofiber mesh, as conventional electrospunscaffolds have been used (Pham Q P, et al., Tissue Eng, 2006; 12:1197-1211; Kumbar S G, et al., Biomed Mater, 2008; 3: 034002;Christopherson G T, et al., Biomaterials, 2009; 30: 556-564; Chua K N,et al., Biomaterials, 2006; 27: 6043-6051).

In an embodiment of the invention, ECFCs cultured on polymer microfibersof the invention exhibit typical membrane expression of endothelialmarkers VEcad and CD31, and cytoplasmic expression of von Willebrandfactor (vWF) (FIG. 7 b-d ). Expression of the endothelial markersdemonstrates that fibrin microfibers support the adhesion and culture ofECFCs. This unique approach allows detailed control of the cellularassembly of microvasculature, as the fibrin microfibers present ablueprint with a unique aligned nanotopography for adhesion of ECFCs.

In embodiments of the invention, ECFCs deposit ECM that wrapscircumferentially around the polymeric microfiber of the invention.Examining the ECFC-deposited ECM organization on the microfiber after 5days in culture, ECFCs deposit ECM proteins, for example, collagen IV,fibronectin and laminin. ECM proteins wrap in discrete circumferentiallyaligned segments on the microfibers, perpendicular to the cellsmacroscopic cellular alignment and intracellular cytoskeletalorganization. This feature of the ECFCs in which they deposit abundantECM (Kusuma S, et al., FASEB J, 2012; 26: 4925-4936), that is assembledcircumferentially on a micro-cylindrical platform, recognizes an activerole of the endothelium in the construction of the extracellularcomponents of the microvasculature. In longer period cultures of ECFCson fibrin microfibers (i.e. >10 days) full coverage of the structures byECM was observed (data not shown). However, at this time point theinitial circumferential organization of the ECM could not be analyzeddue to several layers of ECM being deposited on top of each other. Forquantification purposes, cultures of ECFC were analyzed when ECMorganization was evident, before full coverage was achieved.

Circumferential wrapping of ECM by ECFCs has not been observedpreviously. To elucidate whether the ECFC alignment or the cylindricalstructure and 3D aspect of the scaffold had a direct effect on ECMorganization, two different systems have been used. A flat polymer sheetwith the same nanotopography as the polymer microfibers can be used as afirst scaffold; the flat polymer sheet varies the shape and geometry ofthe scaffold. ECFCs align with the nanotopography of the polymer sheets,but the ECM is deposited with no distinguishable organization. A secondscaffold for use is a polymer microfiber, such as polyethersulfone(PES), coated with a polymer such as fibrin, which maintains themicrofiber's spatial geometry, but has a random surface topography.ECFCs seeded on the polymer microfiber, for example PES fibers, are notinduced to align with the fiber's longitudinal axis, but they depositECM wrapping circumferentially around the polymeric microfiber, similarto ECFCs seeded on fibrin microfibers. Without being bound by theory, itis believed that the cylindrical shape of the fibers, and not thecellular organization induced by the scaffold's nanotopography, isnecessary for ECM circumferential deposition.

The cytoskeleton is known to regulate endothelial alignment (Ranj an A,and Webster T, Nanotechnology, 2009; 20: 305102; Liliensiek S, et al.,Biomaterials, 2010; 31: 5418-5426; Bettinger C J, et al., Adv Mater,2008; 20: 99-103; Lu J, et al., Acta Biomater, 2008; 4: 192-201) anddrive angiogenic responses through ECM-interactions (Bayless K J, DavisG E, Journal of Cell Science, 2002; 115: 1123-1136; Hanjaya-Putra D, etal., J Cell Mol Med, 2009). Cytoskeleton re-arrangement of ECFCs seededon fibrin microfibers through actin and tubulin configuration revealedthat ECFC alignment on the microfibers is not instrumental for thecircumferential deposition of ECM by ECFCs. Inhibition of neither actinfilament nor microtubule polymerization affected ECM circumferentialorganization around the fibrin microfiber. Thus, ECM expression andorganized deposition from ECFCs can be independent of ECFC cellularorganization. Furthermore, shorter culture time periods, for example 3days, which did not always produce a confluent endothelium stillresulted in wrapping ECM, suggesting an independence of cell density onECM organization.

Overall, even though aligned topography of the polymer microfibersinduces ECFC alignment, the findings that either ECFCs seeded on PESfibers or ECFCs seeded on fibrin microfibers with disrupted actin andmicrotubule organization still produce wrapping ECM suggest that ECMorganization is regulated by 3D geometric sensing of curvature, ratherthan by the nano-topography of the scaffold.

In an embodiment of the invention, the microfiber system demonstratedthat circumferential wrapping of the ECM depends on the curvature of themicrofibers. While curvature of nano-scale features of ECM has beensuggested to regulate cellular responses (Vogel V, and Sheetz M, Nat RevMol Cell Biol, 2006; 7: 265-275), its effect on cellular responses atthe micro-scale and during microvascular formation and organization hasnot been previously investigated. In this invention, polymer microfiberswith an aligned nanotopography are generated by electrostretching.Microfibers are produced with uniform and tunable diameters whilepreserving the aligned nanotopography. In one exemplary embodiment,fibrinogen is mixed with alginate in-line and then charged with electricpotential of about 2 kV to about 6 kV; the mixture is extruded through a25-gauge needle. The fibrinogen-alginate solution jet is collected at adistance of about 3 to about 5 cm from the needle tip, in a grounded,rotating bath containing calcium chloride and thrombin as across-linking solution, to generate aligned nanofibers that can later bebundled to form microfibers with an aligned nanotopography. Microfibersare generated with different diameters by varying the collection time,such as from about 5 min and higher, preferably from any time point fromabout 5 min to about 80 min, more preferably from any time point fromabout 7 to about 80 min. Microfiber formation may include crosslinkingnanofibers by any cross-linking method, including ionic crosslinking,ultraviolet crosslinking, enzymatic crosslinking, and chemicalcrosslinking reaction. After formation of crosslinked fibrin-alginatenanofibers, alginate is removed, preferably by soaking fibers in asodium nitrate solution. Excess sodium citrate is washed off and theresulting fibrin nanofibers are collected as an aligned bundle,stretched preferably to 150% of their initial length, and dried.Resulting fibrin microfibers can be wrapped around a custom-made plasticframe and then sterilized for use.

Microfibers with diameter ranging from about 100-500 μm were examined.Microfibers of up to about 450 μm in diameter guide the organizedwrapping of deposited ECM. Preferably, microfibers have a diameter about100 μm to about 450 μm, more preferably about 100 μm to about 400 μm,about 100 to about 350 μm, about 100 to about 300 μm, about 100 to about250 μm, about 200 μm, about 150 μm, and diameters within the ranges,such as about 110 μm, about 120 μm, 130 μm, 140 μm, 150 μm, 200 μm, 250μm, 300 μm, 350 μm, 400 μm, 450 μm. Larger microfibers resulted in amore random ECM organization. This observation is the first to suggestan effect of the microtubular curvature on ECM organization. In contrastto studies using larger diameter templates to study ECM deposition ofperivascular cells (Grassl E, et al., J Biomedical Materials ResearchPart A, 2003; 66: 550-561), the smaller diameter fibers demonstrate therole of curvature at the micro-scale.

The versatility of the polymer microfiber system is evident in allowingsequential and controlled introduction of other cells, which can depositECM. ECFCs will initially have an inverted polarity due to the presenceof the fiber where the luminal surface would be and an absence of atunica media on top. The first step towards correcting this invertedpolarity was obtaining full investment of mural cells on top of theendothelium. Embodiments of the invention relate to the polymermicrofiber cultured with endothelial progenitor cells being furtherseeded with a second cell type, such as perivascular cells, or muralcells. Preferably, the perivascular cell or mural cell is a vascularsmooth muscle cell (vSMC) or pericyte. Both pericytes and vSMCsintroduced independently into the model after ECFC culture on the fibrinmicrofibers were found to attach to the ECFC-seeded microfibers. ThevSMCs and perictyes showed organizations that varied from the random,sporadic attachment typical of muscular venules, to the circumferentialwrapping observed in arterioles under physiological conditions(Standring S, 2008). The organization, however, was in a multilayerconfiguration as opposed to the monolayer formed by the ECFCs, inaccordance to the multilayer organization observed in the tunica mediaof native vessels. The different morphologies observed between vSMCs,pericytes and ECFCs are likely a result of lack of pulsatile flow, whichhas been shown to induce circumferential wrapping of vSMCs in differentstudies (Lee A A, et al., J Biomech Eng, 2002; 124: 37-43; Liu B, etal., Biophysical Journal, 2008; 94: 1497-1507). Indeed, the lack ofinternal pressure in this system is more conducive to the generation ofmuscular venules, which have much lower blood pressures compared toarterioles and contain randomly oriented vSMCs or pericytes (StandringS, 2008).

A further step to correct the polarity of the ECFCs in the model isobtaining a defined lumen to create a hollow microvascular vessel. Afurther advantage of the fibrin microfiber scaffold is itsbiodegradability; fibrin can be easily degraded in a controlled mannerusing plasmin in conditions that do not affect cell viability (Neidert MR, et al., Biomaterials, 2002; 23: 3717-3731). In an embodiment of theinvention, degradation of the fibrin microfibers generates a hollowmicrostructure with a defined lumen, which can be used for applicationsin vivo. Thus, the fibrin microvascular system provides opportunities tocorrect the initial ECFC polarity and to study flow-induced vSMC orpericyte organization post fibrin core degradation.

vSMCs and pericytes attach on ECFC-seeded microfibers and depositextracellular proteins. vSMCs attach and grow on the ECFC layer (FIGS.12 a-c) and deposit collagen Type I and elastin (FIGS. 12 d-g). Thecollagen type I and elastin are located below and in between the vSMClayer, and above the ECFCs (FIGS. 12 e, g). Pericytes attach and grow onthe hydrogel microfiber scaffold (FIGS. 13 a-c), deposit collagen TypeIV (FIGS. 13 d-e), which is located below and in between the pericytelayer and above the ECFCs.

These results establish that co-cultured vSMCs and pericytes depositedECM components that organize the subendothelial connective tissue andinternal elastic lamina, located in between the endothelium and thetunica media, as well as components of the tunica media itself.Co-culture experiments in ECFC media was found to support ECFC,pericytes, and vSMC viability.

Self-supporting structures with the abundant vascular ECM depositionnecessary to withstand pulsatile flow as well as vasoconstriction andvasodilatation are disclosed. ECM proteins such as collagens, laminin,and elastin have been shown to provide these biomechanical properties.To date, the study of ECM protein deposition by different vascular cellsin 3D constructs either alone or in co-culture has been limited (Davis,G. E. and D. R. Senger, Circulation Research, 2005; 97(11): p.1093-1107.). In an embodiment of the invention, increased quantities ofECM were deposited on the 3D microfibers of the invention compared to 2Dcultures after seeding ECFCs, pericytes, and vSMCs in 2D and 3D culture.The 2D surfaces were coated with a thick fibrin hydrogel layer toprovide a similar stiffness and bioactive substrate compared to its 3Dcounterpart. For ECFCs, immunofluorescence microscopy revealed increaseddeposition of ECM proteins Col IV, Fn, and Lmn in 3D compared to 2D(FIGS. 14 a,b). This increased expression was found to be about 2-foldfor Fn and Lmn and almost 4-fold for Col IV, as measured by RT-PCR (FIG.14 c). These results indicate the existence of a geometrical orbiomechanical sensing pathway that upregulates the production of theseproteins comprising the basal lamina of native endothelium (Laurie, G.,et al., Cell Biology, 1982; 95(1): p. 340-344).

Similar studies performed on pericytes and vSMCs revealed these celltypes produce several different ECM proteins that comprise thesubendothelial connective tissue, internal elastic lamina, and ECM ofthe tunica media of blood vessels (Brooke, B. S., et al., Trends in CellBiology, 2003; 13(1):51-56; Hungerford, J. E., et al., DevelopmentalBiology, 1996; 178(2):375-392). In the fibrin hydrogel microfiber of theinvention, both pericytes and vSMCs produced Col types I, III, IV, aswell as Fn and Lmn. Additionally, only vSMCs produced Eln, which is inaccordance to native vasculature where elastic tissue is foundpredominantly in arterioles and arteries with a full vSMC layer and notin smaller capillaries or venules invested only by pericytes (Hibbs, R.G., et al., Am Heart J, 1958. 56(5):662-670; Yen, A. and I. M.Braverman, J Investig Dermatol, 1976. 66(3):131-142; Brooke, B. S., etal., Trends in Cell Biology, 2003; 13(1):51-56). Notably, there is anincrease in average expression of all ECM proteins deposited byperivascular cells in 3D compared to 2D microfibers, though theupregulation was only statistically significant for Col I, Col IV, andEln deposition by vSMCs (FIGS. 15 a-c, 16 a-c). Furthermore, themorphological expression of these proteins was found to be moreorganized in 3D than 2D substrates, and Lmn was found to be depositedextracellularly in 3D microfiber substrates only. This is evidence ofdistinct metabolic pathways that regulate the expression and depositionof different ECM proteins by vSMCs and pericytes. The fibrin microfibersprovide an appropriate microenvironment for abundant, organized ECMdeposition.

In a further embodiment, hollow microvascular vessels are prepared byremoving the polymer microfiber core from the microvascular structures.An advantage of fibrin as a polymer biomaterial, besides its naturalbiocompatibility, bioadhesiveness, and angiogenic promotingcharacteristics (Clark, R. A. F., 2003; 121(5): p. xxi-xxii), is itsbiodegradability in response to enzymes such as plasmin, the enzymeresponsible for eliminating fibrin blot clots in the human body (Rijken,D. C. and H. R. Lijnen, J Thromb Haemost, 2009; 7(1):4-13). In a furtherembodiment, varying plasmin concentrations in serum free media wereshown to degrade fibrin microfibers at different time points (FIG. 17a). The plasmin concentration ranged from 0.1 to 15 CU/mL. In apreferred embodiment, a 12 hr degradation treatment of 0.25 CU/mLplasmin was able to maintain cell viability similar to control cultureconditions while a 24 hr treatment of 0.1 CU/mL plasmin was moredetrimental to cell survival possibly due to the longer period of serumstarvation (FIG. 17 b). Degradation of ECFC-seeded fibrin microfibersafter 5 days of culture revealed that both the 12 and 24 hr treatmentprotocols were able to degrade the fibrin microfiber core whilemaintaining the intact ECM deposited by ECFCs, resulting in an earlymicrovascular structure comprised of an endothelial layer and its basallamina with a clear circular lumen (FIG. 17 c).

In further embodiment of the invention, luminal multicellularmicrovascular structures were created by adding perivascular (mural)cells to constructs that first had been cultured with ECFCs to allowfull endothelial layer formation before introducing the perivascular(mural) cells. Co-cultures were further cultured and shown to becomprised of both ECFCs and mural cells along with their deposited ECM,such as Col III and Col IV for ECFC-pericyte co-cultures (FIG. 12 a).Unexpectedly, ECFC-vSMC co-cultures presented Eln deposition after only5 days of vSMC growth (FIG. 18 b), an elusive achievement in vasculartissue engineering typically shown in vSMC cultures of over four weeksonly (Kim, B. S., et al., Biotechnol Bioeng, 1998, 57(1):46-54; Gao, J.,et al., J Biomedical Materials Research Part A, 2008; 85A(4):1120-1128;Patel, A., et al., Cardiovascular Research, 2006; 71(1):40-49; Long, J.L. and R. T. Tranquillo, Matrix Biol, 2003; 22(4):339-50). Finally, theresulting structures were shown to retain both cellular and ECMformation after degradation of the fibrin core, forming a distinctcircular lumen (FIGS. 18 c, d).

The novel in vitro model system established in accordance with theinvention demonstrates the role of vessel diameter on ECM organizationduring microvascular growth. The system supports a step-wise vascularformation process via introduction of perivascular cells at varying timepoints, a current challenge in microvascular tissue engineering. Thisapproach can be used to develop further a mechanistic understanding ofhuman microvasculature assembly and stabilization in health and disease.

Abbreviations

The following abbreviations may appear in the examples and elsewhere inthe specification and claims:

Actin, beta (ACTB)

aminocaproic acid (ACA)

collagen type I (Col I)

collagen, type I, alpha 1 (COL1A1)

collagen type III (Col III)

collagen, type III, alpha 1 (COL3A1)

collagen type IV (Col IV)

collagen, type IV, alpha 1 (COL4A1)

elastin (Eln)

endothelial cells (ECs)

endothelial colony forming cells (ECFCs)

extracellular matrix (ECM)

fibronectin (Fn)

Glyceraldehyde 3-phosphate dehydrogenase (GAPDH)

laminin (Lmn)

Laminin subunit gamma-1 (LAMC1)

polyethersulfone (PES)

smooth muscle cells (SMCs)

three-dimensional (3D)

two-dimensional (2D)

vascular endothelial growth factor (VEGF)

vascular smooth muscles cells (vSMCs)

EXAMPLES

The following Examples have been included to provide guidance to one ofordinary skill in the art for practicing representative embodiments ofthe presently disclosed subject matter. In light of the presentdisclosure and the general level of skill in the art, those of skill canappreciate that the following Examples are intended to be exemplary onlyand that numerous changes, modifications, and alterations can beemployed without departing from the scope of the presently disclosedsubject matter. The synthetic descriptions and specific examples thatfollow are only intended for the purposes of illustration, and are notto be construed as limiting in any manner to make compounds of thedisclosure by other methods.

Example 1 Preparation and Characterization of Hydrogel Microfibers

To produce alginate hydrogel microfibers, 2-6 kV, 1.5-3.0 wt % alginateand 0.1-0.6 wt % PEG solution were used. The alginate hydrogelmicrofibers were stabilized in a collection bath comprising 20 mM to 100mM CaCl₂) and a rotating collection plate. As a representative example,a starting solution comprising 2.0 wt % alginate (from brown algae,approximately 250 cps viscosity for a 2% solution at 25° C.) and 0.2 wt% poly(ethylene glycol) (PEG, average My ca. 4,000 kDa, Sigma Aldrich,St Louis, Mo.) is used. The starting solution was pumped through a 27gauge needle syringe at 2 mL/hour rate by a syringe pump. A 3-kV voltagewas applied to the needle with a clamp from a high voltage power source(Gamma High Voltage Research, Ormond Beach, Fla.). The rotatingcollection plate in the collection bath has a diameter of about 20 cmand rotates at about 25 rpm. The rotating collection plate waspositioned approximately 4 cm away from the exit of the syringe needle.The 50 mM CaCl₂) solution stabilizes the alginate hydrogel fibers duringcollection. After the hydrogel strings are collected on the rotatingcollection plate, they are allowed to crosslink in the 50 mM CaCl₂)solution for 3 min before use.

With similar flow rates and applied voltages, fibrin, gelatin and HAhydrogel fiber bundles also were prepared. In representative examples,fibrin hydrogel fibers were produced using an aqueous solution thatcontains 0.7 wt % fibrinogen, 1.0 wt % sodium alginate and 0.1 wt % PEG.Upon collection, the hydrogel fibers were crosslinked in 50 mM CaCl₂)with 5 Units/mL thrombin for 20 minutes. If necessary, a higherconcentration of thrombin can be used to shorten the crosslinking time.

As another representative example, aqueous solutions that contain 2.0 wt% fibrinogen from bovine plasma (syringe 1) and 1.5 wt % sodiumalginate/0.2 wt % PEG (syringe 2) were mixed through a Y junction mixerat 1:2 ratio. The mixed solution was pumped through a 25 gauge needlesyringe at 3 mL/hour rate. A 4 kV voltage was used to initiate theelectrostretching process. Upon collection, hydrogel fibers werecrosslinked in CaCl₂)/thrombin solution (50 mM, 5 Units/mL) for 20minutes. After crosslinking, the hydrogel microfibers were soaked in 250mM sodium citrate solution overnight to remove alginate and PEG. Thehydrogel microfibers were then rinsed with distilled water to remove thesodium citrate residue before use.

As an alternative to the protocol disclosed immediately hereinabove,fibrinogen and PEG (1 wt %/0.1 wt %) were directly mixed and processedat 4 kV voltage. The crosslinking solution in this case contained 50 mMCaCl₂) and 20-unit/mL thrombin. After 20 minutes of crosslinking, thehydrogel microfibers can be directly collected for use.

Gelatin hydrogel fibers were prepared with 3.2 wt % methacrylatedgelatin, 0.9 wt % sodium alginate, 0.1 wt % PEG, and 0.4 wt % photoinitiator Irgacure 2959, followed by crosslinking with 50 mM CaCl₂)solution for 5 minutes and then UV-irradiation at λ 365 nm for 10minutes. Methacrylated gelatin was prepared according to a previouslyreported protocol (Nichol et al., 2010). As another representativeexample, gelatin hydrogel fibers were prepared with a solutioncontaining 3.0 wt % methacrylated gelatin, 1.0 wt % sodium alginate,0.15 wt % PEG and 0.4 wt % photo initiator Irgacure 2959 (CIBA SpecialtyChemicals, Basel, CH). Flow rate was set as 2 mL/hour. Other processingparameters were similar to the generation of alginate and fibrindescribed hereinabove. Upon collection, alginate was crosslinked in 50mM CaCl₂) bath for 5 minutes. UV-irradiation at λ 365 nm withMineralight Lamp UVGL-25 (UVP LLC, Upland, CA) was then used tocrosslink the gelatin for 10 minutes. After crosslinking, 250 mM sodiumcitrate solution was used to remove alginate, PEG and photo initiatorovernight. Distilled water was then used to rinse off the sodium citrateresidue before use.

Similarly, HA hydrogel fibers were prepared with 1 wt % thiolated HA,0.7 wt % alginate and 0.2 wt % PEG, and crosslinked with 50 mM CaCl₂)and 1 wt % polyethylene glycol diacrylate (PEGDA). As anotherrepresentative example, HA hydrogel fibers were prepared with 1.0 wt %thiolated HA (Glycosan BioSystems Inc.), 0.75 wt % alginate and 0.2 wt %PEG solution. A 4-kV applied voltage was used to initiate the jet. Theflow rate used was 2 mL/hour. The collection bath contained 50 mM CaCl₂)and 1 wt % PEGDA to crosslink alginate and HA. After 20 minutes ofcrosslinking, alginate, PEG and excess PEGDA can be removed by sodiumcitrate and distilled water, using the protocols for generating alginateand fibrin described hereinabove.

Collagen hydrogel fibers were prepared with 2.0 wt % methacrylatedcollagen (syringe 1) and 1.5 wt % sodium alginate/0.2 wt % PEG/0.4 wt %photo initiator Irgacure 2959 (syringe 2). These solutions were mixedthrough a Y junction mixer at a 1:1 ratio. The combined solution waspumped through a 25 gauge needle syringe at 3 mL/hour rate. 4 kV appliedvoltage was used to initiate the jet. The collection bath contained 50mM CaCl₂). Upon collection, the alginate was crosslinked in the 50 mMCaCl₂) bath for 5 minutes. UV-irradiation at λ 365 nm was then used tocrosslink collagen for 10 minutes. After crosslinking, 250 mM sodiumcitrate solution was used to remove alginate, PEG and photo initiatorovernight. Distilled water was then used to rinse off the sodium citrateresidue before use.

Methacrylated collagen used in this example was prepared by addingmethacrylic anhydride to acid solubilized type I collagen (3 mg/mL, LifeTechnologies, Carlsbad, CA). Before reaction, the collagen solution wasadjusted to pH 7.5 with 0.2 M Na₂HPO4 buffer. Methacrylic anhydride (MA)was added in different ratios to obtain a desired degree crosslinkingcapacity. After an eight hour reaction, the mixture was dialyzed against10 mM HCl for 2 days. Pierce Slide-A-Lyzer Concentrating Solution(Thermo Scientific, Waltham, Mass.) was used to condense the solution tothe desired concentration.

Dehydrated String Bundles.

Dehydrated string bundles known in the art typically are produced by airdrying. Applying an axial stress can exclude the liquid content, speedup the drying process, and enhance alignment. As a representativeexample, fibrin hydrogel strings collected by the protocol disclosedhereinabove was stretched to 160% of its original length and air dried.Alginate hydrogel string collected as described hereinabove also wasstretched to 130% of its original length and air dried. In both cases,the hydrogel strings shrank in diameter and became dehydrated thinstrings. Their average Young's modulus also drastically increased to 10GPa for calcium alginate fibers and 2 GPa for fibrin fibers. Hydrogelstrings of other compositions can be dehydrated in a similar way.Alternatively, hydrogel strings can be frozen and lyophilized to createa porous morphology. Dehydrated strings made by both methods can berehydrated when soaked in water.

Sem Analysis.

Hydrogel microfiber samples were first serially dehydrated in 50%, 60%,70%, 80%, 90%, 95% and 100% ethanol for 15 minutes in each solution,critical point dried, and then sputter-coated with 8-nm thick Au/Pd.Samples were imaged on a JEOL 6700F field-emission SEM (Tokyo, Japan).

Small and Wide Angle X-Ray Scattering.

SAXS experiment was performed at the Cornell High Energy SynchrotronSource (CHESS; Ithaca, NY, USA). Dry or wet hydrogel fibers weresubjected to 10-second exposures of the synchrotron beam (λ=0.11521 nm,beam size: 0.5 mm horizontal×0.1 mm vertical) for 10 times. A 48 mm×48mm 2-D CCD detector with pixel size of 46.9 μm×46.9 μm was used tocollect the scattering data. The averaged intensity readings on eachpixel of the detector were recorded and analyzed with fit2D. WAXSexperiments were performed using a Rigaku R-Axis Spider Diffractometer(Rigaku Americas Corp., The Woodlands, TX, USA) with an image platedetector and a graphite monochromator using Cu Kα radiation (λ=0.15418nm). The instrument was controlled by Rapid/XRD diffractometer controlsoftware (version 2.3.8, Rigaku Americas Corp., The Woodlands, TX, USA).Fibers were grouped into a bundle and secured on the sample stage.Two-dimensional diffraction data were collected for 20 minutes whilerotating the sample stage at 5° per minute. The 2D diffraction data wereradially integrated with 2DP Spider software (version 1.0, RigakuAmericas Corp., The Woodlands, TX, USA).

Mechanical Testing.

Single axial stretching tests were performed over the hydrogelmicrofibers in dry, wet and rehydrated forms with a DMA Q800 unit fromTA Instruments (New Castle, DE, USA). Experiments results revealed that,while dry fibers have limited capacity to elongate (approximately 3-5%strain at break), wet hydrogel fibers were stretched to more than 100%strain before breaking. The average Young's moduli of dry calciumalginate, fibrin, gelatin and HA fibers were 10.0 GPa, 2.2 GPa, 0.8 GPa,and 3.0 MPa, respectively. For wet fibers prior to the drying step, theYoung's moduli were several orders of magnitude lower (717 kPa, 37.3kPa, 2.6 kPa, and 1.3 kPa, respectively). The moduli of rehydratedfibers fell in between the two sets, with 108 MPa, 289 kPa, 4.4 kPa, and58.5 kPa, respectively. In these analyses, sample diameters weredetermined using a light microscope. For wet strings, samples werestretched to break within 30 seconds to minimize the effect of waterevaporation on measurement. The Young's moduli were calculated withinthe initial linear region of the stress-strain curves from the tests.

Example 2 Electromechanical Stretching Setup and Features

This strategy employs electrical and mechanical stretching to inducepolymer chain alignment during spinning of an aqueous polymer solution,followed by rapid chain alignment fixation of the polymer jet viacrosslinking (FIG. 1 a ). The electrostretching setup includes acollection bath comprising a crosslinking solution and a grounded,motor-driven rotating collection plate. The polymer jet is charged witha relatively lower electrical potential of 2-6 kV, including 2, 3, 4, 5,and 6 kV and any fractional value within the range of 2-6 kV, thantypically applied for electrospinning methods known in the art (5-30kV).

In some embodiments, the presently disclosed electromechanicalstretching platform includes a high voltage power supply, a needlesyringe pump, a syringe, and a motor-run rotating metal disc, i.e., thecollection plate. The collection plate can have a diameter of about 20cm. A high voltage DC power is applied to the solution by clamping anelectrode on the syringe needle. The applied voltage is about 2-6 kV,and the flow rate of the solution is about 0.4-4 mL/hour. After the jetis initiated by applying the electrical potential, the stretched stringis collected on the collection plate, which can be positioned about 3 cmto 5 cm away from the exit of the syringe needle in a calcium chloridesolution reservoir, i.e., the collection bath. The angular velocity ofthe collection plate is controlled by a DC motor controller (Dart 15DVE)unit, which was usually operated in a range from about 20 to 80rotations/minute. If more than one solution is involved, such asco-spinning of multiple components or co-axial spinning, an additionalsyringe pump can be used.

In some embodiments, the hydrogel fiber is made from alginate. The goodbiological compatibility and high viscosity of alginate makes it notonly a good candidate to make hydrogel string by itself, but also makesit an ideal template to induce stretching over other materials.

The entire process is conducted in aqueous solutions. For example, asolution of sodium alginate (1.5-3.0 wt %) and polyethylene glycol (PEG,0.1-0.6 wt %) is charged with 2-6 kV positive potential and extrudedthrough a syringe needle at a flow rate of 1-3 mL/h. The PEG in thealginate solution serves as a thickening agent to increase the viscosityof the alginate solution jet (Ji et al., 2006). The electrical fieldcauses the polymer solution to form a jet. The jet is further stretchedupon being collected on the rotating collection plate containing 20-100mM CaCl₂) solution at a collecting distance approximately 3-6 cm fromthe needle tip. The diameter of the collected hydrogel fiber can betuned by adjusting the solution extrusion rate and the angular velocity(20-80 rpm) of the rotating collection plate, which in embodiments wherethe diameter of the rotating collection plate is 20 cm, corresponds to alinear velocity of about 20-84 cm/second. The fiber diameter increaseswith solution extrusion rate and decreases with rotation velocity of therotating collection plate. For example, the average diameter ofindividual calcium alginate fibers can be controlled in the range of 17μm to 116 μm by varying the flow rate of alginate (2 wt %)-PEG (0.2 wt%) solution from 0.7-7 mL/hour at room temperature (FIG. 1 b ). Hydrogelfibers produced with this process have highly uniform diameters (FIG. 1c ), and continuous hydrogel microfibers of any length can be produced.Further, the hydrogel microfibers can be grouped together to formbundles of tunable diameters depending on the number of individualfibers used.

The versatility of the presently disclosed approach can be demonstratedby preparing internally aligned hydrogel microfibers from severalnatural polymers (alginate, fibrin, gelatin (Nichol et al., 2010) andhyaluronic acid (Shu et al., 2004)) using different crosslinking schemes(FIG. 1 c-f ). Calcium alginate hydrogel fibers were initially prepared.One important advantage of the calcium alginate hydrogel system is itsfast gelation rate. Potter et al. have determined the displacement ofthe crosslinking reaction front in 2 wt % alginate solution to beapproximately 20 μm/second in a 50 mM CaCl₂) crosslinking solution, and40 μm/second in 100 mM CaCl₂) (Potter et al., 1994). In theelectrostretching system described above, a 40-μm alginate fiber can beeffectively crosslinked in about 0.5-1.0 second by the CaCl₂) solution.This fast crosslinking scheme allows the incorporation of otherwater-soluble polymers to form polymer blend fibers. Additional enzyme-,UV- or chemical-mediated crosslinking reactions can be used togetherwith calcium ions to further stabilize the hydrogel fibers. In someembodiments, thrombin-mediated crosslinking of fibrinogen can be used toform fibrin-alginate blend fibers, UV light can be used to crosslinkmethacrylated gelatin-alginate fibers, and a Michael-type additionreaction can be used to crosslink thiolated hyaluronic acid-alginatefibers (FIG. 1 d-f ). After crosslinking, alginate and PEG can beremoved from the polymer blend hydrogel fibers by washing the fiberswith sodium citrate. All of these crosslinking and washing steps can becarried out in aqueous buffers under ambient conditions and thereforeare cell-compatible.

Additional components, such as cells and bioactive agents, can beincluded by mixing them with the polymer solution or through anadditional syringe pump. End point mixing nozzle or co-axial stretchingnozzle also can be used. In some embodiments, fibrinogen is mixed withalginate and the mixture is co-crosslinked with CaCl₂) and thrombin. Inother embodiments, gelatin and hyaluronic acid strings are formed bymixing methacrylated gelatin or acrylated hyaluronic acid with alginateand co-crosslinking with CaCl₂), photo initiator Irgacure 2959 and UVlight. In these embodiments, alginate is used as a template material andcan be dissolved in calcium sequester solution, such as sodium citrate,EDTA or even PBS solution for fast or slow alginate removal.

Electrostretched hydrogel microfiber bundles are mechanically strongerand easier to handle than the typical hydrogels of the same compositionand size. As a demonstration, an electrostretched calcium alginatehydrogel fiber bundle was used to lift a 10-g metal weight (FIG. 1 i ),and also two individual alginate gel strings were tied into a micro-knotusing forceps (FIG. 1 j ). On the contrary, bulk alginate hydrogels oralginate fibers prepared with the same concentration of alginate, butwithout employing the presently disclosed stretching process cannotwithstand such manipulations. Due to the improved mechanical propertyand ease of handling, such hydrogel materials can be further fabricatedinto other forms like films, tubes and more (FIGS. 1 l-m ).

To probe the structural origin for enhanced mechanical properties,birefringence imaging of the electrostretched hydrogel fibers wasconducted. The extinction of light at the cross-point of fibers (FIG. 1k ) indicated strong polymer chain alignment within the hydrogelmicrofiber bundles.

Example 3 Scanning Electron Microscope Images and Small Angle X-RayScattering (SAXS) Patterns

The alignment structure was further confirmed on critical point-driedhydrogel microfiber bundles utilizing scanning electron microscope (SEM;FIGS. 2 a-2 i ). As shown in FIG. 2 a-c , fibrin, methacrylated gelatin,and thiolated HA hydrogels prepared using simple mixing and crosslinkingsteps formed random nanofiber mesh networks. In contrast, theelectrostretched hydrogel microfibers exhibited preferential alignmentalong the fiber axis (FIG. 2 d-f ). Such highly porous and alignedsurface texture also is very different from recently developed fibrinmicrothreads, which are dense and smooth on the surface (Cornwell andPins, 2007; Grasman et al., 2012). Grouping of individual fibers intobundles followed by further stretching (usually 30-100% of the initiallength) and dehydration resulted in dense microfibers with alignedgrooves and surface textures (FIG. 2 g-h ). These dry fibers can berehydrated to about 50-100% of their original diameter depending ontheir drying processes.

To confirm molecular alignment within the hydrogel fibers, both in dryand hydrated forms, their small angle X-ray scattering (SAXS) patternswere analyzed (FIG. 3 a-b ), which showed strong anisotropic scatteringprofiles, indicating preferential orientation along the fiber axis. As acomparison, both dry and hydrated non-stretched alginate samples gave anisotropic scattering pattern (FIG. 3 c ). The wide-angle X-rayscattering (WAXS) analysis of dry calcium alginate microfibers alsoshowed a reflection profile that was indicative of an oriented polymercrystalline phase (FIG. 3 d ). The reflection pattern also confirmedthat the polymer chains are oriented preferentially along the microfiberaxis indicated by the arrow. Results with fibrin and gelatin strings aresimilar to that of calcium alginate (FIG. 4 ).

The preferential alignment of polymer chains within the microfibersgreatly improved mechanical properties of the hydrogel fibers. FIG. 3e-g shows the Young's moduli of dry, wet, and hydrated hydrogel fiberbundles. While dry fibers have limited capacity to elongate(approximately 3-5% strain at break), wet hydrogel fibers were stretchedto more than 100% strain before breaking. The average Young's moduli ofdry calcium alginate, fibrin, gelatin and HA fibers were 10.0 GPa, 2.2GPa, 0.8 GPa, and 3.0 MPa, respectively. For wet fibers prior to thedrying step, the Young's moduli were several orders of magnitude lower(717 kPa, 37.3 kPa, 2.6 kPa, and 1.3 kPa, respectively). On the otherhand, the moduli of rehydrated fibers fell in between the two sets, with108 MPa, 289 kPa, 4.4 kPa, and 58.5 kPa, respectively. According to thetheory proposed by MacKintosh et al., the modulus of an entanglednetwork scales with concentration (MacKintosh et al., 1995). Therefore,the modulus and stiffness of the hydrogel fibers can be further adjustedby varying the concentration of starting materials, such as alginate andfibrinogen, and crosslinking density. As substrate modulus plays animportant role in regulating cellular behaviors like proliferation,migration and differentiation, the ability to tune hydrogel fibermodulus and stiffness over such a wide range makes the presentlydisclosed hydrogel fiber matrix versatile for a wide range ofapplications (Engler et al., 2006; Discher et al., 2005). These analysesconfirm that hydrogel fibers prepared by electrostretching exhibitpolymer chain alignment along the microfiber axis. In contrast, hydrogelsamples prepared by simple extrusion have an isotropic structure.

The relatively low electrical potential applied to the polymer solution(a measured current of 4-6 μA), the aqueous solvent, and ambientcrosslinking conditions make this process compatible with cellencapsulation. In some embodiments, alginate is not a favorable cellscaffold matrix due to the lack of cell adhesion moieties. Therefore, insome embodiments, fibrin, gelatin or hyaluronic acid are blended in tothe hydrogel fibers, each employing a unique second crosslinking step tofurther stabilize the hydrogel fiber matrix. For example, a solution offibrinogen, alginate, and PEG can be mixed with cells, and subjected tothe electrostretching condition as described above.

Hydrogel fibers are rapidly crosslinked by the calcium solution in thecollection bath, followed by crosslinking of fibrinogen into fibrinnetwork with thrombin. Similarly, methylated gelatin and thiolated HAcan be used instead of fibrinogen using the corresponding crosslinkingmethods discussed in FIG. 1 (detailed conditions are listed in Table 1;all fibers describe in Table 1 were spun at 3-5 kV electrical potentialand collected on a rotating collection plate spun at 20-80 rpm). Allthese crosslinking methods are cell-compatible. After the secondcrosslinking step, alginate and PEG can be removed with sodium citrate,if a higher degree of porosity is desired.

TABLE 1 Spinning Parameters for different Hydrogel Microfibers HydrogelSpinning Solution Crosslink Composition Concentration (wt %) CrosslinkMethod Condition Alginate 0.75-3.0% Alginate Ionic 25-100 mM CaCl₂0.1-0.4% PEG crosslinking Fibrin + 0.67-2.0% Fibrin Enzymatic and ionic5 U/mL thrombin Alginate 0.25-2.5% Alginate crosslinking 50 mM CaCl₂0.1-0.2% PEG Gelatin + 1.0-3.2% Gelatin Ionic & UV- 50 mM CaCl₂ Alginate0.24-0.86% Alginate crosslinking 0.50% Irgacure 0.40-1.0% Irgacure 10min UV at 365 nm Hyaluronic Acid + 1.0-2.0% HA Ionic & 1% PEGDA &Alginate 1.5% Alginate Michael addition 50 mM CaCl₂ 0.1-0.4% PEGcrosslinking Fibrin 0.67-2.0% Fibrin Enzymatic 20 U/mL thrombin 0.1-0.2%PEG crosslinking Collagen + 0.67-2.0% Collagen Ionic & UV- 50 mM CaCl2Alginate 0.25-2.5% Alginate crosslinking 0.50% Irgacure 0.1-0.2% PEG 10min UV at 365 nm

Example 4 Loading Drugs or Other Bioactive Agents in HydrogelMicrofibers

The hydrogel microfibers can be loaded with drugs or other active agentsin situ or through post-loading. In situ loading can be achieved byincluding these active agents in the polymer solution. As an example,nanoparticles or growth factors can be directly added into alginatesolution at various concentrations and processed using the typicalprocess for making the hydrogel microfibers in Example 1. Thecrosslinked calcium alginate network thus encapsulates the nanoparticlesand growth factors inside the microfiber.

Alternatively, microfibers can be loaded with drugs or other bioactiveagents by soaking either the hydrated or dehydrated microfiber in asolution containing the drugs or active agents. In one example ofpost-loading, alginate/fibrin blend microfibers are soaked in a solutioncontaining growth factors at various concentrations for 12 hours, andthen dried in the air.

Drugs and bioactive agents include, but not limiting to, grow factors,small molecular weight compounds that can promote cell growth, enhancecell differentiation and maturation, or facilitate cell migration andtissue organization. In some specific examples, bioactive agents areselected from glial cell-derived neurotrophic factor, nerve growthfactor, bone morphogenic factor, hepatic growth factor, vascularendothelial growth factor, and the like.

Example 5 Culture Mammalian Cells with Hydrogel Microfibers

Cell Encapsulation into Hydrogel Microfibers.

Mammalian cells, including but not limited to adipose tissue-derived.stem cells, Schwann cells, oligodendrocytes, etc. can be encapsulatedinto fibrin or fibrin-alginate hydrogel microfibers at a density of2,000-10,000 cells/μL. To produce such cell-laden hydrogel microfibers,cells can be suspended in fibrinogen solution, which is then mixed at1:2 volume ratio through a. syringe with a solution of 1.5 wt % sodiumalginate and 0.2 wt % PEG solution, and charged at +4 kV potential, andelectrostrectched according to the procedure described in Example 2. Thecollection bath will contain a stabilization solution of 50 mM CaCl₂, 5units/mL thrombin and 5% glucose to maintain the physiologicalosmolarity. The crosslinking step can be conducted similarly as thatdescribed in Example 2. After crosslinking, cell-laden fibers can becollected, transferred into Petri dish with media, and cultured in 5%CO) incubator at 37° C.

Culture Cells on the Surface of Hydrogel Fibers.

Microfibers prepared by the method described here can be used as amicro-scaffold for cell culture. Mammalian cells can be seeded onto thesurface of hydrogel microfibers and cultured using standard cell culturetechniques.

Example 6 Discussion

The presently disclosed subject matter demonstrates that a high degreeof axial alignment of polymer chains inside hydrogel microfibers can beinduced by a combination of electrical and mechanical stretchingeffects. Polymer chain alignment induction during the electrospinningprocess has been previously reported (Catalani et al., 2007; Bellan andCraighead, 2008; Fennessey and Farris, 2004). For example, PEG polymerchain can be aligned when the PEG solution is subjected to a 14-kVelectric potential (Kakade et al., 2007). This observed alignment arisesbecause the PEG chain has flexible C—O ether bonds in the backbone,facilitating the PEG chain alignment along water molecule dipoleorientation in response to the strong electrical field (Kakade et al.,2007). This method may not be applicable to other polymer-solventsystems, however, particularly for the presently disclosed biopolymers,which exhibit longer relaxation times and coiled chain conformations.

In the presently disclosed methods, these barriers are circumvented byreducing the electric potential and extending the jet stretching time.In some embodiments, the average air travel time of the alginatesolution jet ranged from about 100 msec to about 500 msec before it iscollected on a rotating collection plate in a collection bath containingthe crosslinking solution, in contrast to a typical air travel timeabout 10 msec for the liquid jet in electrospinning before solventevaporation step (Reneker, 2000). Again, without wishing to be bound toany one particular theory, it is thought that this extended jetstretching time likely allows alginate chains to align better with theelectric field.

In further embodiments, this electric field-induced polymer chainalignment can be further enhanced by mechanical stretching (FIG. 5 ).According to theoretical models, a high degree of polymer chainalignment can be achieved during the uniaxial stretching of a polymersolution jet if the Weissenberg number, defined as the product of strainrate {acute over (ε)} and the conformational relaxation time λ, isgreater than 1 (Larson and Mead, 1993). Despite the high strain rates(10⁵-10⁶ s⁻¹) commonly observed in polymer solution jets duringelectrospinning, high degrees of alignment are usually difficult toachieve. This difficulty is likely due to the rapid solidification ofelectrospun fibers under typical spinning conditions, which does notgive sufficient time for polymer chains to align (Inai et al., 2005;Zong et al., 2002). In some embodiments, under the presently disclosedelectrostretching conditions, the alginate solution is collected withoutsignificant solvent evaporation, and the solution jet has an estimatedstrain rate {acute over (ε)} of 10-70 s⁻¹. Although the strain rate isnot very high, it is compensated by the long relaxation time due to thehigh molecular weight of alginate and PEG. Thus, the mechanical shearinduced by the rotating collection plate can significantly contribute tothe higher degree of alignment of the polymer chain. This analysis issupported by the observation that a faster rotating velocity leads tofibers with higher tensile modulus, indicating a higher degree ofalignment enhanced by stronger mechanical stretching (FIG. 6 ).

The Young's modulus of wet fibers increased with the angular velocity ofthe rotating collection plate. These results suggest that a higherdegree of mechanical stretching—under a higher angular velocity of therotating collection plate—induces a higher degree of polymer chainalignment, manifested by a higher Young's modulus.

Without wishing to be bound to any one particular theory, it is thoughtthat the efficacy of the presently disclosed method also relies on aneffective crosslinking or fixation of the induced polymer chainalignment. The presently disclosed subject matter provides differentcrosslinking strategies that are applicable for a wide selection ofbiopolymers. These crosslinking methods also are complementary so thatit is possible to prepare blended fibers having different compositionsto afford multi-functionalities—a feature particularly suitable forregulating cell adhesion, tissue compatibility, permeability, andsurface conjugation of ligands.

Further, the biodegradability of the presently disclosed fibers can betailored by blending polymers with different degrees of sensitivity tohydrolysis and degradative enzymes. In some embodiments, the degradationcan be triggered on demand. In some other embodiments, alginate fiberscan be dissolved by treating the fibers with sodium citrate solution. Infurther embodiments, fibrin fibers can be degraded by plasmin and HAfibers can be degraded by hyaluronidase.

In some embodiments, the entire fiber spinning and crosslinking processis conducted in aqueous solutions under ambient conditions, making itamenable to cell encapsulation inside hydrogel fibers. In particular,the low electric potential (2-6 kV) in contrast to electrospinning andlow mechanical shear ensure high cell viability in hydrogel fibers.

In some embodiments, the hydrogel fibers generated by the presentlydisclosed electrostretching method exhibit excellent mechanicalproperties while maintaining sufficient porosity and water content(>90%, usually 98%-99%). This characteristic is in contrast withhydrogel fibers prepared by simple extrusion, in which case polymersolutions are pressed through a small orifice and crosslinked during orafter extrusion. Hydrogel fibers produced by the extrusion method do notexhibit a high degree of polymer chain alignment (FIG. 3 c ), and aretherefore mechanically weaker and more challenging to handle thanhydrogel fibers produced by electrostretching. Although this limitationin extruded fibers can be overcome by lowering the water content andincreasing crosslinking density, these denser fibers are not suitablefor cell encapsulation. The porosity of the electrostretched hydrogelfibers also can be easily tuned by varying the input polymerconcentration, composition and crosslinking density.

Accordingly, the presently disclosed subject matter provides methods togenerate hydrogel microfibers with a high degree of polymer chainalignment induced by a combination of electrical and mechanicalstretching, and facilitated by the long polymer chain relaxation time,and effective crosslinking schemes. Using this concept, internallyaligned hydrogel microfiber bundles of calcium alginate, fibrin,gelatin, hyaluronic acid, collagen and their blends have been produced.These microfibers exhibit enhanced mechanical properties as a result ofthe polymer chain alignment while maintaining high water content andporosity. The facile preparation conditions are conducive to cellencapsulation in generating “cellular strings.” Due to theirbiodegradable nature and unique geometry and surface topography, theyare ideal scaffold candidates for generating aligned tissue structuresThe development of these new hydrogel fibers represents an importantstep toward successful fabrication of hierarchically organized cellularstructures in 3D.

Examples of Microvascular Structures

The invention can be further understood in view of the followingnon-limiting examples.

Cell Culture

Unless indicated otherwise, the following cells and culture conditionswere used in the experiments. Human ECFCs (Lonza, Walkersville, MD) wereused for experiments between passages 5 and 9. ECFCs were expanded inflasks coated with type I collagen (BD Biosciences, Franklin Lakes, NJ)in Endothelial Basal Medium-2 (EBM-2; Lonza) supplemented with EGM-2Bulletkit (Lonza) and 10% fetal bovine serum (FBS; Hyclone, Logan, UT).ECFCs were fed every other day, passaged every 5 to 7 days with 0.05%trypsin/0.1% ethylenediaminetetraacetic acid (EDTA; Invitrogen,Carlsbad, CA). Human vSMCs (ATCC, Manassas, VA) were used betweenpassages 4 and 9 and cultured in F-12K medium (ATCC) supplemented with0.01 mg/ml insulin (Akron Biotech, Boca Raton, FL), 10% FBS (Hyclone),0.05 mg/ml ascorbic acid, 0.01 mg/ml transferrin, 10 ng/ml sodiumselenite, 0.03 mg/ml endothelial cell growth supplement, 10 mM HEPES,and 10 mM TES (all from Sigma-Aldrich, St. Louis, MO). Human placentalpericytes (Promocell, Heidelberg, Germany) were cultured in PericyteGrowth Media (Promocell) and used between passages 7 and 10. Media waschanged every other day and cells were passaged every 5 to 7 days with0.05% trypsin.

Immunofluorescence Staining and Confocal Microscopy Imaging

Cell-microfiber constructs were fixed with 3.7% formaldehyde (FisherChemical, Fairlawn, NJ) for 15 min, permeabilized with 0.1% Triton X-100solution (Sigma-Aldrich) in 3.7% formaldehyde for 10 min, washed threetimes with PBS, and incubated for 1 h at room temperature with theindicated primary antibodies (Table 1). After rinsing with PBS threetimes, samples were then incubated with the appropriate secondaryantibodies or conjugated phalloidin (Table 1) at room temperature for 1h. Samples were then rinsed with PBS three times, and counterstainedwith DAPI for 10 min. Z-stack and cross-sectional images were obtainedand processed using confocal microscopy (LSM 510 Meta, Carl Zeiss Inc.,Thornwood, NY). Epifluorescence images were obtained using an Olympus®BX60 microscope.

TABLE 1 Antibodies Used in the Studies Reagent Type Name Host VendorDilution Microtubule Nocodazol NA Sigma-Aldrich 3.3 μM disrupting agentActin Cytochalasin D NA Sigma-Aldrich   1 ug/mL disrupting agent PrimaryCD31 mouse Dako 1:100 Antibody vWF mouse Dako 1:100 VEcad mouse SantaCruz 1:100 Biotechnology SM22 rabbit Abcam 1:200 Collagen I mouse Abcam1:100 Collagen III mouse Santa Cruz 1:100 Biotechnology Collagen IVrabbit Santa Cruz 1:100 Biotechnology Elastin mouse Abcam 1:100Fibronectin rabbit Sigma-Aldrich 1:100 Laminin rabbit Abcam 1:100Conjugated DAPI NA Roche 1:1000 Antibody Diagnostics Alexa Fluor shroomInvitrogen 1:50 488 Phalloidin Secondary Alexa Fluor 488 goat Invitrogen1:500 antibody anti rabbit IgG Alexa Fluor 546 donkey Invitrogen 1:500anti mouse IgG Alexa Fluor 546 donkey Invitrogen 1:500 anti rabbit IgGAlexa Fluor 647 donkey Invitrogen 1:500 anti rabbit IgG

Transmission Electron Microscopy

Samples were prepared for transmission electron microscopy (TEM)analysis as described previously (Hanjaya-Putra et al., Blood, 2011;118: 804-815). Briefly, samples were fixed with 3.7% formaldehyde, 1.5%glutaraldehyde in 0.1 M sodium cacodylate, 5 mM CaCl₂), and 2.5% sucroseat room temperature for 1 h and washed 3 times in 0.1 M cacodylate/2.5%sucrose (pH 7.4) for 15 min each. The cells were post-fixed withPalade's OsO₄ on ice for 1 h, en bloc stained with Kellenberger uranylacetate overnight, dehydrated through a graded series of ethanol, andthen embedded in EPON™ epoxy resin. Sections of 80 nm were cut, mountedonto copper grids, post-stained in 2% uranyl acetate and Reynolds leadcitrate, and viewed using a Phillips® EM 420 transmission electronmicroscope (FEI). Images were captured with an Olympus® Soft ImagingSystems Megaview III CCD digital camera.

Scanning Electron Microscopy

Hydrogel microfiber samples were first serially dehydrated in 50%, 60%,70%, 80%, 90%, 95% and 100% ethanol for 15 min in each solution,critical point dried, and then sputter-coated with 8 nm thick Au/Pd(gold/palladium particles). Samples were imaged on a field-emissionscanning electron microscopy (SEM) (JEOL 6700F, Tokyo, Japan).

Image and Statistical Analyses

The cell areas and perimeters were measured by fitting an ellipse toeach cell using the LSM 510 software. Cytoskeletal alignment angle wascalculated by measuring the angle between the long axis of each ellipseand the longitudinal axis of the microfiber, found by drawing a line atthe edges of the microfiber in its image projection. ECM angle oforientation was measured using the LSM 510 software by drawing a linefollowing the ECM deposition and finding the angle between the line andthe longitudinal axis of the microfiber. Graphs were plotted with 5-95%confidence intervals. Unpaired two-tailed Welch-corrected t-tests wereperformed where appropriate (GraphPad Prism® 5.01, GraphPad Software,San Diego, CA). Significance levels were determined between samplesexamined and were set at *p<0.05, **p<0.01, and ***p<0.001.

Example 7 ECFC Attachment and Alignment on Fibrin Microfibers

A new approach to create aligned hydrogel microfibers using anelectrostretching process of polymer materials is disclosed. Uniquecharacteristics of the electrostretched hydrogel microfibers are theinternal and topographical alignment of the fibrous structure, generatedas a result of both electrical field and mechanical shear-inducedpolymer chain alignment as described above. Furthermore, the diameter ofa microfiber is controlled and uniform as a result of the bundling andprocessing of the individual nanofibers that compose the hydrogelmicrofibers. Fibrin gels have been extensively used to studymicrovasculature assembly (Dickinson L E, et al., Soft Matter, 2010; 6:5109-5119; Bayless, K J, and Davis, G E, Biochemical and BiophysicalResearch Communications, 2003; 312: 903-913; Davis G E, and Bayless K J,Microcirculation, 2003; 10: 27-44; Bayless K J, et al., RGD-DependentAmerican Journal of Pathology, 2000; 156: 1673-1683; Dickinson L E, etal., Lab Chip, 2012; 12: 4244-4248), vSMC responses (Ahmann K A, et al.,Tissue Eng Part A, 2010; 16: 3261-3270; Long J L, and Tranquillo R T,Matrix Biol, 2003; 22: 339-350) and multicellular organization (LesmanA, et al., Biomaterials, 2011; 32: 7856-7869). Fibrin is used as thematrix material to prepare hydrogel microfibers as a template for thestep-wise creation of microvasculature of the invention.

Preparation of 3D Fibrin Hydrogel Microfiber

Fibrin hydrogel microfibers were generated by the electrostretchingmethod (FIG. 1A). An aqueous solution of 1.5 wt % alginate(Sigma-Aldrich) was in-line mixed with 2 wt % fibrinogen (Sigma-Aldrich)at feeding rates (flow rates) of 2 ml/h and 1 ml/h, respectively. Bothsolutions were dissolved in 0.2 wt % poly(ethylene oxide) (PEO)(Mw=4,000,000, Sigma-Aldrich) prior to spinning. The mixed solution wasthen charged with 4 kV electric potential and extruded through a25-gauge needle. The fibrinogen-alginate solution jet was collected, ata distance of about 3-5 cm from the needle tip, in a grounded, rotatingbath (20 cm diameter, 20-40 rotation/min) containing 50 mM CaCl₂solution with 5 units/ml thrombin (Sigma-Aldrich) as a cross-linkingsolution (FIG. 7 ). After spinning, fibers were left in the collectionsolution for 15 min. To generate microfibers with different diameters,collection times were varied from 7-80 min, including 7, 10, 15, 17, 20,22, 26, 27, 35, 40, 45, 50, 55, 60, 70, 75, and 80 min. The crosslinkedfibrin-alginate fibers were then soaked in 0.2 M sodium citrateovernight to remove calcium ions and dissolve alginate. Fibers weresoaked in water for 30 min to remove sodium citrate, stretched manuallyto about 150% of their initial length by placing the ends of themicrofiber on supports such as pipettes and extending the supports tofor example 150% of their original distance, and air-dried for 30 min.Fibers were wrapped around a custom-made plastic frame, then sterilizedby soaking in 75% ethanol for 2 min followed by rinsing twice withsterile water. Microfiber diameter was measured from confocal Z-stackprojections.

Cell Seeding and Culture on Fibrin Hydrogel Microfibers

ECFCs were seeded on microfibers of 15-cm length total at a density of4×10⁵ cells/ml in 5 ml of ECFC media supplemented with 50 ng/mL of VEGF(Pierce, Rockford, IL, USA). The seeding tube was continuously rotatedon a tumbler (Labquake, Dubuque, Iowa) for 24 h at 37° C. to facilitatecell attachment. Frames with ECFC seeded microfibers were thentransferred to 35 mm Petri dishes using tweezers, and cultured in thesame media in a CO₂ incubator at 37° C. Media was refreshed every otherday thereafter.

Results

Using the electrostretching method, fibrin hydrogel microfibersexhibited longitudinally-aligned nanotopography (FIG. 7 a ). This is animportant feature as sub-micron (<1 μm, but greater than 100 nm) scaletopographic features have been shown to increase EC adhesion, migrationand orientation (Ranjan A, and Webster T, Nanotechnology, 2009; 20:305102; Liliensiek S, et al., Biomaterials, 2010; 31: 5418-5426;Bettinger C J, et al., Adv Mater, 2008; 20: 99-103; Lu J, et al., ActaBiomater, 2008; 4: 192-201).

The seeding of ECFCs was facilitated by continuous rotation (FIG. 7 a ).After 24 hrs, ECFCs attached to the microfibers throughout the surface(FIG. 7 e ). Within 5 days in culture, ECFCs were found to be elongatedand aligned longitudinally with the microfibers as indicated by F-actinstaining (FIG. 7 b-d ). ECFCs covered the microfiber surfacecontinuously, and exhibited typical membrane expression of endothelialmarkers VEcad and CD31 (FIG. 7 b, 7 d ), and cytoplasmic expression ofvon Willebrand factor (vWF) (FIG. 7 c ), demonstrating that fibrinmicrofibers support the adhesion and culture of ECFCs.

Example 8 ECM Deposition from ECFCs on Fibrin Microfibers

While the importance of ECM deposition in vascular development has beenrecognized, few studies have looked at ECM production by theendothelium. ECFCs deposit collagen IV, fibronectin and laminin in anorganized web-like structure when cultured on Petri dishes (Kusuma S, etal., FASEB J, 2012; 26: 4925-4936). To establish a reliable in vitromodel of microvasculature, we characterized the ECM protein depositionby the endothelium on hydrogel microfibers. ECFCs were seeded on fibrinhydrogel microfibers as described in Example 7.

ECFC seeded on fibrin microfibers deposited laminin, collagen IV, andfibronectin after one day in culture (FIG. 8 a ). On Day 5, ECFCscompletely covered the fibrin microfiber, and abundant ECM depositionwas observed (FIG. 8 b ). In contrast to what was observed previously onPetri-dishes (Kusuma S, et al., FASEB J, 2012; 26: 4925-4936), the ECMproteins deposited by ECFCs on hydrogel microfibers were organized. ECMproteins laminin, collagen IV, and fibronectin wrapped around themicrofibers, perpendicular to the EC orientation, along the microfiber'scircumference, as observed in FIGS. 8 c, 8 f, and 8 g . This is incontrast to the web-like structure observed on Petri-dishes.Specifically, on fibrin microfibers, the individual collagen IVnano-fibrils also seem to follow this macroscopic circumferentialalignment at the nano-scale, with fibrils being deposited next to eachother in an aligned manner to create a ribbon of circumferentiallyaligned collagen (FIG. 8 c, 8 g ). Further analysis revealed this wasalso true for microfibers of sizes up to about 370 μm, and that thelargest microfibers tested (about 445 μm) did not have a distinguishablecollagen nano-fibril orientation (FIG. 8 d-f ). On Petri-dishes, ECMnano-fibrils were deposited without any apparent alignment, in aweb-like structure formation (Kusuma S, et al., FASEB J, 2012; 26:4925-4936). Moreover, these ECM structures appeared to be distributedeither below or among the ECFCs (FIG. 8 d-e ), resembling basal laminaorganization found in native microvessels. It should be noted thatsimilarly to what we observed on Petri-dishes (Kusuma S, et al., FASEBJ, 2012; 26: 4925-4936), ECFCs did not express or deposit Collagen I(data not shown).

Example 9 ECM Deposition from ECFCs on Fibrin Sheets and PES Fibers

It was previously demonstrated that line-grating topography influencesEC adhesion, alignment, and elongation (Ranjan A, and Webster T,Nanotechnology, 2009; 20: 305102; Liliensiek S, et al., Biomaterials,2010; 31: 5418-5426; Bettinger C J, et al., Adv Mater, 2008; 20: 99-103;Lu J, et al., Acta Biomater, 2008; 4: 192-201). To probe if the alignednanotopography on fibrin microfibers is responsible for ECFC alignmentand coordinated deposition of laminin, collagen IV, and fibronectin, wefirst examined their deposition on flat (2D) fibrin sheets with similaraligned nanotopography (FIG. 9 e ) as the fibrin hydrogel microfibers byvarying the dimensionality and cylindrical shape of the scaffold.

Preparation of 2D Fibrin Nanofiber Sheets

Fibrin-alginate hydrogel nanofibers were prepared according to the sameelectrostretching method as described in Example 1 above until thecollection step in a rotating bath. The collected fibrin-alginatehydrogel nanofibers were then wrapped around a modified plastic frame toform a sheet of hydrogel nanofibers while slightly stretching thenanofibers to ensure proper alignment. The fibrin nanofiber sheets wereplaced in a 0.2 M sodium citrate solution overnight to remove alginate,followed by a 30 min wash in water to remove excess sodium citrate.Fibrin nanofiber sheets were then sterilized with 75% ethanol and rinsedtwice with sterile water. The resulting nanofiber sheets are distinctfrom microfibers as these are not bundled, stretched, and air-dried toform a microfiber, but instead are collected from the rotating bath in a2D nanofiber sheet formation.

Cell Seeding and Culture on 2D Fibrin Sheets

Cells were seeded by placing 5×10⁵ ECFCs in a concentrated solution ofcells (about 2×10⁶ cells/ml) directly on top of the fibrin nanofibersheet. After 5 min the cell solution that had filtered through thesheets was collected and reseeded on top of the fibrin nanofiber sheets.This process was repeated 3 times, after which the same culture media asused for 3D fibers was added to the samples before they were placed in ahumidified incubator at 37° C. in a 5% CO₂ atmosphere. Media wasrefreshed every other day up to 5 days of culture.

Preparation of 3D Polyethersulfone Fibers

Solid polymer fibers were prepared as a control according to a modifiedelectrospinning protocol. In brief, polyethersulfone (PES) (GoodfellowCambridge Limited, UK, Mw 55,000) was dissolved in 30 wt % DMSO andelectrospun under an electric potential of 5 kV. The feed rate of PESsolution was 12 ml/h to initiate a polymer jet, which was collected in agrounded, rotating ethanol bath (20-40 rotations/min) to extract thesolvent. The collection distance was set to 5 cm. After 10 min inethanol, PES strings were removed from the bath and air-dried.

After electrospinning, PES fibers were wrapped around a seeding framesimilarly to the fibrin hydrogel microfibers. Samples were thenplasma-treated for 5 min before soaking for 5 min in a 10 units/mlthrombin in 15 mM CaCl₂ solution. Thrombin-coated PES fibers were thenimmersed in a 0.2% fibrinogen solution diluted in 0.9% NaCl forfibrinogen polymerization into fibrin. Excess fibrin coating on theframe and outside of the fibers was removed before sterilization with75% ethanol for 1-2 min. Samples were rinsed twice with sterile water,after which cell seeding was performed similarly to the fibrinmicrofibers as described in Example 1.

Results

ECFCs seeded on fibrin nanofiber sheets were effectively aligned withthe nanotopography. However, the collagen IV, fibronectin and lamininproduced by ECFCs exhibited a random organization, as opposed to theperpendicular orientation with respect to cell alignment observed in the3D microfibers (FIG. 9 a ). This result indicates that the microfibergeometry may be crucial to the specific organization of ECM moleculessecreted by ECFCs.

Similarly as observed in FIG. 8 , ECFCs completely covered the PESmicrofibers and deposited ECM molecules after 5 days of culture (FIG. 9b ). While ECFCs grown on the PES microfibers did not necessarily have arandom orientation on the PES fibers, and in fact often exhibited apartial diagonal orientation (FIG. 9 c ), the ECFC-deposited ECMproteins were found to wrap around the PES microfiber in a similarmanner to ECFC-deposited ECM on fibrin microfibers, as evidenced bymeasuring the angles between ECM ribbons and the fiber's longitudinalaxis (FIG. 9 d ). Note that the average angle in both cases is close to90 degrees, demonstrating perpendicular alignment, and that thedistribution (represented by the height of the boxes and also shown asstandard deviation in FIG. 9 e ) is small, signifying most ECM ribbonshad an orientation close to 90 degrees.

Electrospun PES microfibers coated with fibrin used to generatemicrofibers maintain the dimensionality and geometry of the fibrinmicrofibers but with a random nanotopography (FIG. 9 f ). Before coatingthe PES fibers with fibrin, the surface of PES fibers is smooth (FIG. 9g (ii)), however, an uncoated PES fiber is not bioadhesive, and ECFCattachment after seeding is not detected (data not shown). Furthermore,PES fiber does not present the same bioactive substrate to the ECFCs asthe fibrin fibers. Therefore, the PES fibers were coated with fibrin,resulting in the random, non-aligned nano-topography of coatingpresented in FIG. 9 g (i). Such PES microfibers have a similar diameter(240±45 μm; data not shown) as fibrin microfibers and thus enablesinvestigation of whether the uniaxial alignment topography contributesto the unique cellular activity and ECM organization.

Example 10 ECM Organization: Dependence on ECFC Alignment andMicrotubule Organization

Actin and microtubule disruption studies were used to determine whetherECFC actin filament alignment and microtubule organization through actinand tubulin configuration directs ECM organization. Cytochalasin D is anactin destabilizing agent, and nocodazole is a microtubulepolymerization disturbing agent.

Cell Seeding and Culture

ECFCs were seeded on fibrin microfibers as described in Example 1 above.

Actin and Microtubule Disruption Studies

Cytochalasin D or nocodazole were dissolved in DMSO (Table 1). ECFCswere cultured on Petri dishes (control) or fibrin microfibers in ECFCmedia with 50 ng/mL of VEGF (Pierce, Rockford, IL, USA) supplementedwith either 1 g/mL cytochalasin D or 3.3 M nocodazole, from either day 0or day 1 after seeding. F-actin or α-tubulin organization and ECMdeposition was analyzed after 1, 2, or 3 days of treatment. Finalconcentration of DMSO in cell culture medium was kept at 0.1% (v/v).Controls were treated with DMSO alone at the same concentration.

Results

For Cytochalasin D added to the culture media at 24 h after ECFC seedingon microfibers, forty-eight hours after treatment the unique wrappingarrangement of the deposited ECM molecules around the fibrin microfibersstill present after 24 and 48 hrs of treatment (FIG. 10 a and FIG. 10 f). No difference was found in the average angle of ECM orientation or inthe variance of all angles measured (FIG. 10 d-e ). It should be notedthat ECM deposition was observed also in control treatments of ECFCs inPetri dishes (FIG. 10 g ) and that similar organization of wrapping ECMwas observed when cytochalasin D was applied through the entire 3 daysof culture (FIG. 10 h ).

Likewise, when nocodazole, a microtubule polymerization disturbingagent, was added to the culture media at 24 h after ECFC seeding onmicrofibers, a similar ECM wrapping pattern was observed (FIG. 10 b ;FIG. 10 i ) even though microtubule formation was disrupted compared tosamples with no drug treatment (FIG. 10 c ). Similarly, no differencewas found in the average angle of ECM orientation or in the variance ofall angles measured (FIG. 10 d-e ). In addition, ECM deposition was alsoobserved in the 2D control group with the same treatment (FIG. 10 j )),and similar organization of wrapping ECM was observed when Nocodazole Dwas applied through the entire 3 days of culture (FIG. 10 k ).

Overall, while treatment of ECFCs seeded on the nanopatternedmicrofibers with either cytochalasin D or nocodazole effectively alteredactin and microtubule organization, respectively, it did not alter thewrapping organization of the deposited ECM molecules as compared tocontrol cultures (FIG. 10 ). Also, even after only 3 days of culturewhen the endothelium layer was not always confluent and therefore thecell density was lower, ECM organization was still found to becircumferential (FIG. 10 c-e , FIG. 10 l-m ).

Example 11 ECM Organization: Dependence on Microfiber Curvature

As indicated, the ECM organization is independent of the cytoskeletonorganization of the ECFCs, but is influenced by the geometry of themicrotubular structure. Here the diameter of the tubular structure tomodulate the ECM organization was evaluated.

Preparation of Fibrin Hydrogel Microfiber

Hydrogel microfibers were prepared as in Example 1 above. Microfibers ofdifferent sizes were prepared by varying the collection time of theelectrostretching process, thus changing the number of nanofibers ineach microfiber bundle. Fibrin microfibers with an average diameter of107.1±11.7 μm, 136.1±12.1 μm, 372.0±27.3 μm, and 443.4±30.6 μm wereprepared. These microfibers were processed similarly and thus exhibitedsimilar nanotopographical alignment, inducing alignment of ECFCs withthe longitudinal axis of the microfibers on all sizes (data not shown).

Results

While seeded-ECFCs were confluent after 5 days in culture on allmicrofibers, decreasing organization of the ECM molecules was apparentas the fiber diameter increased (FIG. 11 a ). Measuring the anglesbetween ECM ribbons and the microfiber's longitudinal axis revealed thatmicrofibers with diameters smaller than about 400 μm had average anglesclose to 90° with a small distribution, signifying perpendicularorientation. On the largest diameter tested (avg. 452.1±26.7 μm), weobserved a non-circumferential ECM organization as evidenced by asignificant increase in the distribution of the angles between ECMribbons and the fiber's longitudinal axis (FIG. 11 b ). Since the anglevalues measured range from 0° to 180°, a perfectly random distributionof angles would average to the mean of 90° as well. However, a samplewith ECM wrapping would have most of these values close to 90°, having asmall variance compared to a sample presenting random ECM orientation.Indeed, the largest microfibers resulted in ECM angles with a markedlyhigher standard deviation compared to the smaller microfibers (FIG. 11 c).

Also, when collagen IV has a wrapping organization, the individualcollagen nano-fibrils also follow this macroscopic circumferentialalignment at the nano-scale (FIG. 8 c, 8 g ). This was also true formicrofibers of about 105 μm to about 370 μm, and the largest microfiberstested (about 445 μm) did not have a distinguishable collagennano-fibril orientation (FIG. 11 d-f ).

Example 12 vSMCs and Pericyte Attachment and New ECM Deposition onECFC-Seeded Microfibers

An advantage of the new fibrin microfiber system is the opportunity toco-culture mural cells, for example, vSMCs and pericytes, to study theirinteractions with the endothelial layer as well as the deposition of ECMcomponent that compose the tunica media (Jain R K, Nat Med, 2003; 9:685-693; Carmeliet P, Nature, 2000; 407: 249-257). Here, ECFC-seededfibrin microfibers are co-cultured with vSMCs and/or pericytes toevaluate the effect on organization and ECM deposition.

Cell Seeding and Culture

ECFC-seeded fibrin hydrogel microfibers were prepared as described inExample 1 above.

Human vSMCs (ATCC, Manassas, VA) were used between passages 4 and 9 andcultured in F-12K medium (ATCC) supplemented with 0.01 mg/ml insulin(Akron Biotech, Boca Raton, FL), 10% FBS (Hyclone), 0.05 mg/ml ascorbicacid, 0.01 mg/ml transferrin, 10 ng/ml sodium selenite, 0.03 mg/mlendothelial cell growth supplement, 10 mM HEPES buffer, and 10 mM TESbuffer (all from Sigma-Aldrich, St. Louis, MO).

vSMCs were seeded on 5-7 day ECFC-seeded fibrin microfibers at 1-4×10⁵cells/ml in 5 ml of 0.5% serum or regular ECFC media, tumbled for 24hours, and then transferred to 35 mm Petri dishes to continue culture.Media was changed every other day thereafter.

Human pericytes (PromoCell, Heidelberg, Germany) were used betweenpassages 5 and 9 and cultured in Pericyte Growth Media (PromoCell)according to manufacturer's instructions. Pericytes were seeded on 5 dayECFC-seeded fibrin microfibers at 4×10⁵ cells/ml in 5 ml of ECFC mediasupplemented with 30 mM aminocaproic acid (to prevent fibrinolysis),tumbled for 24 hours, and then transferred to 35 mm Petri dishes tocontinue culture for 9 more days (10 days of co-culture total). Mediawas changed every other day.

Results

vSMCs seeded on ECFC-coated fibrin microfibers attached and grew on theECFC layer, forming a bilayer cellular construct (FIG. 12 a ).Occasionally, vSMCs were found to have a random orientation on thestructures (FIG. 12 a ), and in some instances they wrapped around themicrofibers (FIG. 12 b ). However, more often they aligned with thelongitudinal axis of the microfibers (FIG. 12 c ). vSMCs cultured for 3and 5 days deposited collagen type I and elastin (FIG. 12 d-g ), locatedbeneath the vSMC layer and above SM22 negative cells, the ECFCs (FIG. 12e, g ).

Pericytes cultured for 10 days on ECFC-seeded fibrin microfibersattached and grew on the microfiber scaffold, forming a multilayercellular construct (FIG. 13 ). Pericytes either wrapped or aligned withthe longitudinal axis of the microfibers (FIGS. 13 a-c ). Pericytesdeposited collagen type IV, located beneath and in between the pericytelayer, and above the ECFCs (FIGS. 13 d-e ).

Example 13 3D Multicellular Microvascular Structure

Following the successful culture of different vascular cell types,namely ECFCs, pericytes, and vSMCs on fibrin microfibers, increased ECMprotein production by the cells was investigated. Also, the fibrin coreof the microfiber was degraded leaving both cellular and ECM formationintact, resulting in multicellular microvascular structures with adistinct circular lumen that recapitulate the multilayer organizationfound in native blood vessels.

Cell Culture

All cells were cultured in a humidified incubator at 37° C. and 5% CO₂.Human ECFCs (Lonza, Walkersville, MD) were cultured on collagen I (BDBiosciences, Franklin Lakes, NJ) coated flasks in Endothelial BasalMedium-2 (EBM-2; Lonza) supplemented with EGM-2 Bulletkit (Lonza) and10% fetal bovine serum (FBS; Hyclone, Logan, UT) and used forexperiments between passages 7 and 10. Media was changed every other dayand cells were passaged every 5 to 7 days with 0.05% trypsin(Invitrogen, Carlsbad, CA).

Human vSMCs (ATCC, Manassas, VA) were used between passages 7 and 10 andcultured in F-12K medium (ATCC) supplemented with 0.01 mg/ml insulin(Akron Biotech, Boca Raton, FL), 10% FBS (Hyclone), 0.05 mg/ml ascorbicacid, 0.01 mg/ml transferrin, 10 ng/ml sodium selenite, 0.03 mg/mlendothelial cell growth supplement, 10 mM HEPES, and 10 mM TES (all fromSigma-Aldrich, St. Louis, MO). Media was changed every third day andcells were passaged every 5 to 7 days with 0.25% trypsin (Invitrogen).

Human placental pericytes (Promocell, Heidelberg, Germany) were culturedin Pericyte Growth Media (Promocell) and used between passages 7 and 10.Media was changed every other day and cells were passaged every 5 to 7days with 0.05% trypsin.

Preparation of 3D Fibrin Hydrogel Microfiber

Fibrin hydrogel microfibers were generated by the electrostretchingmethod described in Example 7. Briefly, 1.5 wt % alginate(Sigma-Aldrich) was mixed in-line with 2 wt % fibrinogen (Sigma-Aldrich)at flow rates of 2 ml/h and 1 ml/h, respectively. Both solutions weredissolved in 0.2 wt % polyethylene oxide) (PEO) (Mw=4,000,000,Sigma-Aldrich) prior to electrospinning. A 4 kV electric potential wasapplied to the solution before extrusion through a 25-gauge needle. Thesolution jet was collected in a grounded, rotating bath (30-45rotation/min) containing 50 mM CaCl₂ solution with 10 units/ml thrombin(Sigma-Aldrich) for 35 min. Fibers were left in the collecting solutionfor 10 min and then soaked overnight in 0.25 M sodium citrate todissolve the alginate. Fibers were then soaked in water for 60 min,bundled and stretched to 150% of their initial length, and air-dried for60 min. Microfibers were wrapped around a custom-made plastic frame andsterilized by soaking in 75% ethanol for 2 min followed by rinsing twicewith sterile water.

Cell Seeding and Culture on Fibrin Hydrogel Fibers

ECFCs, vSMCs, and pericytes were seeded on fibers as previouslydescribed in Examples 1 and 6. Briefly, cells were seeded on microfiberswrapped on a frame at a density of 4×10⁵ cells/ml in 5 ml of ECFC mediaand tumbled overnight at 37° C. to facilitate cell attachment. ForECFCs, media was supplemented with 50 ng/mL of vascular endothelialgrowth factor (VEGF; Pierce, Rockford, IL, USA), whereas for pericytesmedia was supplemented with 30 mM aminocaproic acid (ACA;Sigma-Aldrich). At day 2 frames were transferred to 35 mm Petri dishesand cultured in the same media in a 5% CO₂ humidified incubator at 37°C. Media was changed every other day thereafter. For co-cultures, vSMCsor pericytes were seeded on 5 day ECFC-seeded fibrin fibers at 4×10⁵cells/ml in 5 ml of ECFC media or in ECFC media with 30 mM ACA, tumbledfor 24 hours, and then transferred to 35 mm Petri dishes to continueculture. Media was changed every other day thereafter.

Preparation of 2D Fibrin Coated Surfaces

Cell culture 6-well plates and coverslips were coated with a solution of0.2% and 0.1% fibrinogen in normal saline, respectively, and thencrosslinked with 10 U/mL thrombin in 15 mM CaCl₂. The solutions wereincubated for 15 min before being sterilized with 75% ethanol for 2 minand rinsed twice with water.

Cell Seeding and Culture on 2D Fibrin Surfaces

Cells were seeded at 1×10⁵ on 6-well plates and at 5×10⁴ on coverslipsin the same culture media as their 3D counterpart. Samples were placedin a humidified incubator at 37° C. in a 5% CO₂ atmosphere and media wasrefreshed every other day up to 5 days of culture.

Plasmin Treatment

Fibrin microfibers with or without cells were treated with 15, 1, 0.25,and 0.1 CU/mL plasmin from human plasma (Athens Research and Technology,Athens, GA) in DMEM (Life Technologies, Grand Island, NY) for 1, 6, 12,and 24 hrs respectively in a humidified incubator at 37° C. in a 5% CO₂atmosphere. Samples were imaged immediately after treatment.

Live/Dead Assay

ECFCs cultured in 2D and treated with plasmin were incubated with 2 μMcalcein AM and 4 μM ethidium homodimer (Invitrogen) in PBS for 30 min at37° C. and 5% CO₂. Samples were imaged immediately after and the numberof live and dead cells was counted using ImageJ (NIH, Bethesda, MD).

Immunofluorescence Staining and Imaging

Samples were processed as previously described above.

Reverse Transcription Polymerase Chain Reaction

Two-step RT-PCR was performed on 2D and 3D constructs as previouslydescribed (Wanjare, M., S. Kusuma, and S. Gerecht, Stem Cell Reports,2014; 2(5): 561-575). Briefly, total RNA was extracted using a TRIzolprotocol (Gibco, Invitrogen) and quantified using an ultravioletspectrophotometer. RNA was transcribed at 1 μg per sample using reversetranscriptase MMLV (Promega Co., Madison, WI) and oligo(dT) primers(Promega) according to manufacturer's instructions. TaqMan Universal PCRMaster Mix and Gene Expression Assay (Applied Biosystems, Foster City,CA) were used to quantitate the expression of COL1A1, COL3A1, COL4A1,FN1, ELN, LAMC1, ACTB, GAPDH, and 18S genes (Life Technologies). The PCRstep was performed for 40 cycles of 15 seconds at 95° C. and 1 min at60° C. in an Applied Biosystems StepOne Real-TimePCR System (AppliedBiosystems). Relative expressions of the genes were normalized to theamount of ACTB, GAPDH, or 18S in the same cDNA by using the comparativeΔΔC_(T) method provided by the manufacturer. Samples were run intriplicate.

Statistical Analyses

All experiments were performed in triplicate for at least 2 biologicalreplicates. Paired t-tests were performed to compare significancebetween 2D and 3D gene expression (GraphPad Prism 5.01, GraphPadSoftware, San Diego, CA) and graphs were plotted with SEM. Significancelevels were determined between samples examined and were set at *p<0.05,**p<0.01, and ***p<0.001.

Results

Deposition of ECM Proteins by ECFCs in 3D Vs 2D Substrates

We had originally shown the ability of ECFCs to deposit basal laminaproteins Col IV, Fn, and Lmn when cultured in two-dimensional (2D) petridishes (Kusuma S, et al., FASEB J, 2012; 26: 4925-4936). We alsodemonstrated in Example 5 above the importance of micro-scale curvaturein regulating the organized deposition of these ECM proteins by ECFCsutilizing fibrin microfibers of different sizes. Specifically, we showedthat microfibers with diameters ranging from 100 to 400 μm guidecircumferential ECM deposition by ECFCs. Here, the 3D fibrin microfibers(average diameter 188.2±13.9 μm) upregulate the amount of ECM proteinsproduced by ECFCs. As shown in FIG. 14 , ECFCs deposit wrapping Col IV,Fn, and Lmn after 5 days in culture on 3D fibrin microfibers and 2Dfibrin-coated substrates. However, higher amounts of all ECM proteinswere observed in 3D vs 2D (FIG. 14A-B). Quantitative RT-PCR performed onthese substrates also revealed a higher expression of the genes encodingthese ECM proteins in 3D than in 2D (FIG. 14C). Together, these resultsshow that the fibrin microfibers promote not only proper ECMorganization, but also increase ECM deposition compared to 2D cultures.

Deposition of ECM Proteins by Pericytes in 3D Vs 2D Substrates

Regulation of the quantity of ECM proteins deposited by perivascularcells by substrate dimensionality was shown with pericytes seeded onfibrin microfibers following the same protocol used for ECFCs. Notably,pericytes were capable of degrading fibrin microfibers before 5 days, afinding in line with the demonstrated fibrinolytic effects of othermural cells (Grassl, E. D., et al., J. Biomedical Materials Res., 2002.60(4):607-612) but contradictory to studies that have found pericytes toexpress fibrinolysis inhibitors (Kim, J. A., et al., Brain endothelialhemostasis regulation by pericytes. J Cereb Blood Flow Metab, 2005.26(2): 209-217). To allow study of ECM deposition after 5 days inculture the media was supplemented with 30 mM ACA, which has beenpreviously used to inhibit plasmin activity 9 (Ahmann, K. A., et al.,Tissue Eng Part A, 2010. 16(10): 3261-70; and Anonick, P. K., et al.,Arterioscler Thromb, 1992. 12(6):708-716.

As shown in FIG. 15A, pericytes attach and grow on fibrin microfibers.Similarly to ECFCs, pericytes also produce Col IV, Fn, and Lmn both in2D and 3D. Additionally, they deposit Col I and Col III (FIG. 15A-B).Eln deposition was not observed neither in 2D nor 3D cultures asconfirmed by RT-PCR (FIG. 15C). However, the amount of ECM deposited bypericytes was not distinguishably different in 2D vs 3D based onconfocal microscopy. Quantitative RT-PCR analysis revealed there is anincreased expression of COL1A1, COL3A1, COL4A1, FN1, and ELN, but thatthis increase is variable and not statistically significant (FIG. 15C).

ECM deposition in 2D appears either randomly organized (FIG. 15B I andIII), non-polymerized (FIG. 15B II and V), or following local pericyteorientation (FIG. 15B IV), whereas in 3D ECM deposition is organizedparallel to the cell's orientation, which follows the longitudinal axisof the microfiber (FIG. 15A). Additionally, Lmn shows a polymerizedextracellular deposition in 3D compared to 2D (FIG. 15A V, 15B V, and15D). Lastly, pericytes can grow in a multilayer organization on themicrofibers (FIG. 15D), as opposed to the monolayer formed by ECFCs.

Deposition of ECM Proteins by vSMCs in 3D Vs 2D Substrates

Previously in Example 6 we demonstrated that vSMCs can be introduce onECFC-seeded fibrin microfibers and obtain full mural cell investment onthe developing microvascular structures. Additionally, vSMC ECM proteinsEln and Col 1 were deposited between the ECFC and vSMC layer (Example6). Here, the three-dimensionality of the fibrin microfibers induces ahigher ECM deposition by vSMCs than 2D cultures. As seen in FIG. 16A,SMCs can attach and grow directly on the microfibers.

Furthermore, vSMCs deposit Col I, III, IV, Eln, Fn, and Lmn both in 2Dand 3D cultures (FIG. 16A-C). Remarkably, Col I can present a wrappingorientation similar to the previously demonstrated deposition of Col IVby ECFCs when cultured in 3D, compared to a sparse mostly intracellularexpression in 2D (FIG. 10AI and 10BI). Additionally, Eln, Fn, and Lmncan follow an aligned deposition with cellular orientation along thelongitudinal axis of the microfiber in 3D (FIG. 16A IV-VI). In contrast,Fn and Lmn presented a random or intracellular expression in 2D,respectively (FIG. 16B V-VI).

Regarding ECM quantity, Col IV and Eln were upregulated in 3D cultures,with Eln deposition being almost null in 2D but uniformly deposited in3D (FIG. 16A IV and 16B IV). Indeed, all proteins were found to beexpressed in higher quantities in 3D than in 2D via RT-PCR, thoughCOL1A1, COL4A1, and ELN were the only genes significantly upregulatedwith average fold differences of 3.7, 3.0, and 4.9 compared to 2D,respectively (FIG. 16C). Similarly to pericytes, vSMCs in 3D grow in amultilayer organization and deposit polymerized Lmn extracellularly(FIG. 16D).

Fibrin Microfiber Degradation

The fibrin microfiber core was degraded after endothelial cell (EC)layer formation while maintaining both cellular viability and intact ECMorganization. Fibrin microfibers without cells were treated withdifferent concentrations of plasmin. Plasmin effectively degraded themicrofibers in a concentration dependent manner; microfibers treatedwith a range of concentrations for 24 hrs revealed that 15, 1, 0.25, and0.1 CU/mL plasmin degraded the microfibers in about 1, 6, 12, and 24hrs, respectfully (FIG. 17A). Fetal bovine serum present in regularculture media was found to interfere with the degradation process (datanot shown), and therefore all plasmin treatments were performed inserum-free media.

Degradation treatment maintained ECFC viability after treating confluentlayers of ECFCs with the same conditions found for 1, 6, 12, and 24 hrdegradation times. While both 1 hr and 6 hr treatments resulted in poorcell viability (data not shown), the 12 hr and 24 hr treatments resultedin 80.6±4.3% and 65.2±1.8% live cells, respectively (FIG. 17B), comparedto 90.7±7.4% when cells were cultured in regular media (not shown).

We then tested the 12 and 24 hr degradation treatments on ECFC-seededfibrin microfibers that had been in culture for 5 days. As shown in FIG.17C, the previously demonstrated circumferential organization of ECMproteins Col IV, Fn, and Lmn was maintained after microfiber degradationin both 12 hr and 24 hr treatment conditions (FIG. 17C I and IIrespectively). Insert in FIG. 17C II shows a cross-sectional view,demonstrating the resulting lumen.

Luminal Multicellular Microvascular Structure

Multicellular microvascular structures were prepared by culturing ECFCson fibrin microfibers for 5 days, introducing pericytes and/or vSMCs ontop of the endothelial monolayer, and continuing culture for 5 moredays. As shown in FIG. 18A-B, the resulting structures contain both anendothelial monolayer expressing the tight junction protein vascularendothelial cadherin (VEcad) and a fully invested perivascularmulticellular layer expressing vSMC and pericyte marker SM22. Moreover,abundant ECM protein deposition was observed, including ECM proteinsdeposited by both ECFCs and mural cells as shown above (Col IV and Fn;FIGS. 18A III and 18B III), mural cells only (Col III; FIG. 18A III),and vSMCs only (Eln, FIG. 18B III). The resulting structures had acell-ECM wall thickness of 19.9±3.1 μm and 13.6±0.6 μm for pericyte andvSMC co-cultures, respectively.

Constructs were treated with 0.25 CU/mL plasmin for 12 hrs andstructures analyzed for lumen formation. As shown in the orthogonalviews and 3D reconstructions of confocal microscopy images in FIG.18C-D, both pericyte and vSMC co-cultures resulted in a distinctcircular lumen comprised by ECs and perivascular cells, as evidenced byVEcad and SM22 expression. Furthermore, the structures maintained theECM protein composition, as shown for Col III and IV (FIG. 18C) and Fnand Eln (FIG. 18D). Orthogonal projections are shown for top Z-stackslices in FIG. 18C and FIG. 18D. The 3D reconstructions show an angleview of half of the resulting cylindrical microvascular structure.

Due to confocal microscopy limitations when imaging large multicellularstructures only one halve of the structure is visible (the side closestto the objective), resulting in a semi-circular cross-section. Sampleswere therefore flipped over and imaged on both sides to confirmstructure uniformity, and images presented are representative images forboth sides of the structures.

In describing the present invention and its various embodiments,specific terminology is employed for the sake of clarity. However, theinvention is not intended to be limited to the specific terminology soselected. A person skilled in the relevant art will recognize that otherequivalent components can be employed and other methods developedwithout departing from the broad concepts of the current invention. Allreferences cited anywhere in this specification are incorporated byreference as if each had been individually incorporated.

REFERENCES

All publications, patent applications, patents, and other referencesmentioned in the specification are indicative of the level of thoseskilled in the art to which the presently disclosed subject matterpertains. All publications, patent applications, patents, and otherreferences are herein incorporated by reference to the same extent as ifeach individual publication, patent application, patent, and otherreference was specifically and individually indicated to be incorporatedby reference. It will be understood that, although a number of patentapplications, patents, and other references are referred to herein, suchreference does not constitute an admission that any of these documentsforms part of the common general knowledge in the art.

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Although the foregoing subject matter has been described in some detailby way of illustration and example for purposes of clarity ofunderstanding, it will be understood by those skilled in the art thatcertain changes and modifications can be practiced within the scope ofthe appended claims.

That which is claimed:
 1. A biodegradable microfiber having alongitudinally aligned nanotopography comprising biodegradable,electro-mechanically stretched, hydrogel, polymers, wherein alignment ofthe polymers is stabilized during gelation of the hydrogel.
 2. Themicrofiber of claim 1, wherein the biodegradable, electro-mechanicallystretched, hydrogel polymers are in the form of nanofibers, wherein thenanofibers are parallel to each other.
 3. The microfiber of claim 2,wherein the nanofibers are substantially free of a ceramic.
 4. Themicrofiber of claim 2, wherein the nanofibers form a conduit with adiameter in the range of 20 micrometers to 20 mm.
 5. The microfiber ofclaim 2, comprising a solid bundle of the nanofibers with a diameter of0.1 to 100 nm.
 6. The microfiber of claim 1, wherein the hydrogelpolymers have a water content of greater than about 90%.
 7. Themicrofiber of claim 1 having a diameter from about 100 μm to about 500μm based on the outer circumference of the microfiber.
 8. The microfiberof claim 1, wherein the polymers are selected from the group consistingof alginate, fibrin (fibrinogen), gelatin, hyaluronic acid, and acombination thereof.
 9. The microfiber of claim 8, wherein the polymersare fibrin polymers.
 10. The microfiber of claim 1, further comprisingendothelial progenitor cells seeded on the polymer microfiber.
 11. Themicrofiber of claim 10, wherein the endothelial progenitor cells areendothelial colony forming cells.
 12. The microfiber of claim 11,wherein the endothelial colony forming cells are aligned longitudinallyto the microfiber.
 13. The microfiber of claim 12, wherein theendothelial colony forming cells deposit extracellular matrix proteins,and wherein the extracellular matrix proteins are circumferentiallyorganized, wrapping around the microfiber.
 14. The microfiber of claim13, wherein the extracellular matrix proteins include laminin, collagenIV, and fibronectin.
 15. The microfiber of claim 14, wherein collagenIV, laminin, and fibronectin are deposited in higher quantities on themicrofiber than on 2D cultures.
 16. The microfiber of claim 1, furthercomprising perivascular cells seeded on the polymer microfiber.
 17. Themicrofiber of claim 16, wherein the perivascular cells are pericytes.18. The microfiber of claim 17, wherein the pericytes depositextracellular matrix proteins, and wherein the extracellular matrixproteins are longitudinally organized along the microfiber.
 19. Themicrofiber of claim 18, wherein the extracellular matrix proteinsinclude collagen types I, III, IV, laminin, and fibronectin.
 20. Themicrofiber of claim 19, wherein collagen types I, III, IV, laminin, andfibronectin are deposited in higher quantities on the microfiber than on2D cultures.
 21. The microfiber of claim 16, wherein the perivascularcells are vascular smooth muscle cells.
 22. The microfiber of claim 21,wherein the vascular smooth muscle cells deposit extracellular matrixproteins, and wherein the extracellular matrix proteins arelongitudinally, randomly, or circumferentially organized along themicrofiber.
 23. The microfiber of claim 22, wherein the extracellularmatrix proteins include collagen types I, III, IV, elastin, laminin, andfibronectin.
 24. The microfiber of claim 23, wherein collagen types I,III, IV, elastin, laminin, and fibronectin are deposited in higherquantities on the microfiber than on 2D cultures.
 25. The microfiber ofclaim 10, further comprising a second cell type seeded on the fibrinmicrofiber.
 26. The microfiber of claim 25, wherein the second cell typeis a mural cell.
 27. The microfiber of claim 26, wherein the mural cellis vascular smooth muscle cell or a pericyte.
 28. The microfiber ofclaim 27, wherein the vascular smooth muscle cell or the pericyteencircles, is randomly oriented, or is longitudinally oriented withrespect to the fibrin microfiber.
 29. The microfiber of claim 28,wherein the vascular smooth muscle cell deposits collagen type I andelastin, and the pericyte deposits collagen type IV.
 30. Thebiodegradable microfiber of claim 13 wherein the extracellular matrixproteins are induced to circumferentially organize and wrap around themicrofiber by the longitudinally aligned nanotopography of themicrofiber wherein the microfiber is substantially free of a chemicalthat promotes cell alignment.
 31. A biodegradable, electro-mechanicallystretched, hydrogel polymer nanofiber, wherein the polymer is stabilizedduring gelation of the hydrogel.
 32. The nanofiber of claim 31comprising a longitudinally aligned nanotopography.
 33. The nanofiber ofclaim 31 wherein the nanofiber is substantially free of a ceramic. 34.The nanofiber of claim 31, wherein the hydrogel polymer nanofiber has awater content of greater than about 95%.